Eosinophils do not drive acute muscle pathology in the mdx mouse model of Duchenne muscular dystrophy

Abstract

Eosinophils are present in muscle lesions associated with Duchenne Muscular Dystrophy (DMD) and dystrophin-deficient mdx mice that phenocopy this disorder. While it has been hypothesized that eosinophils promote characteristic inflammatory muscle damage, this has not been fully examined. Here, we generated mice with the dystrophin mutation introduced into PHIL, a strain with a transgene that directs lineage-specific eosinophil ablation. We also explored the impact of eosinophil overabundance on dystrophinopathy by introducing the dystrophin mutation into IL5-transgenic (IL5tg) mice. We evaluated the degree of eosinophil infiltration in association with myofiber size distribution, centralized nuclei, serum creatine kinase and quantitative histopathology scores. Among our findings: (1) eosinophils were prominent in quadriceps muscles of 4-week-old male mdx mice but (2) no profound differences were observed in quantitative measures of muscle damage when comparing mdx vs. mdx.PHIL vs. mdx.IL5tg mice, despite dramatic differences in eosinophil infiltration (CD45⁺CD11c⁻Gr1⁻MHCII^loSiglecF⁺ eosinophils at 1.2 ± 0.34% vs. < 0.1% vs. 20 ± 7.6% of total cells, respectively). Further evaluation revealed elevated levels of eosinophil chemoatttractants eotaxin-1 and RANTES in muscle tissue of all three dystrophin-deficient strains; eotaxin-1 concentration in muscle correlated inversely with age. Cytokines interleukin-4 and interleukin-1 receptor antagonist were also detected in association with eosinophils in muscle. Taken together, our findings challenge the long-held perception of eosinophils as cytotoxic in dystrophin-deficient muscle; we show clearly that eosinophil infiltration is not a driving force behind acute muscle damage in the mdx mouse strain. Ongoing studies will focus on the functional properties of eosinophils in this unique microenvironment.

Introduction

Duchenne Muscular Dystrophy (DMD) is a progressive disorder of muscle function that affects 1 in 3500 to 5000 male children worldwide [1]. DMD results from mutations in the X-linked gene encoding dystrophin, a 427 kDa protein that links the cytoskeleton to the extracellular matrix. At the cellular level, the absence of dystrophin destabilizes mechanical forces and signal transduction, leading to myofiber degeneration and subsequent inflammatory responses. Ultimately, this leads to inability to walk, diaphragm lesions and cardiomyopathy [2–4]. Current treatment for DMD is largely supportive [5, 6], although therapies directed at in vivo repair of genetic lesions hold promise [7–9].

Several mouse strains have been used to model primary muscle lesions characteristic of DMD [10–12]. Among these, the DMD^mdx (mdx) mouse has been the subject of the most intense study. This strain, which has a nonsense/stop codon in exon 23 of the dystrophin gene, arose spontaneously in C57BL/10 mice [10]. Although muscles from mdx mice contain little to no dystrophin, the mice experience a mild form of DMD due to compensatory expression of muscle utrophin [12]. However, as with human subjects with DMD, skeletal muscle in mdx mice undergo degeneration and regeneration; acute lesions have been detected in skeletal muscle of mdx mice at 3–4 weeks of age with primary regeneration observed several weeks thereafter [13].

Several groups have explored these lesions in detail and have suggested that the inflammatory response is a critical component of acute muscle damage [14–16]. Among these findings, eosinophilic leukocytes have been detected in muscle lesions in both human subjects with DMD and mdx mice [17–20]. Prednisone, used therapeutically to maintain ambulation in patients with DMD, resulted in diminished numbers of eosinophils in muscle lesions when administered to mdx mice [17, 18]. Likewise, eosinophil major basic protein (MBP-1), while not specifically cytotoxic in vivo, activated splenocytes and promoted muscle fibrosis in in vitro assays [19].

Here, we asked whether eosinophils were instrumental in promoting acute muscle lesions characteristic of dystrophin deficiency. To accomplish this goal, we generated mice in which the dystrophin mutation (mdx) was introduced into the PHIL strain [21] which are mice that maintain a transgene that directs lineage-specific eosinophil ablation during hematopoiesis. We also explored the impact of eosinophil overabundance in these lesions by generating crosses between mdx mice and those carrying the transgene that directs T-cell specific overexpression of interleukin-5 [22]. With these new mouse lines, we examined the impact of eosinophils in acute muscle lesions characteristic of dystrophin-deficiency.

Materials and Methods

Mice.

Mdx (C57BL/10ScSn-Dmd^mdx/J, 001801) and wild-type C57BL/10 (C57BL/10ScSnJ, 000476) mice were purchased from the Jackson Laboratory (Bar Harbor, MD). IL5tg/NJ.1638 mice [22] and PHIL mice [21] were provided by the Lee Laboratories, the latter courtesy of Dr. Elizabeth Jacobsen. The mdx.IL5tg and mdx.PHIL strains were generated by crossing female mice homozygous for the mdx mutation with male IL5tg mice; expression of the IL5 transgene was verified by determining serum IL-5 levels using the mouse IL5 Duoset ELISA (DY405, R&D Systems). Presence of the PHIL transgene was confirmed by genotyping as described [21]. All mouse strains were maintained as in-house colonies in the 14BS vivarium on-campus at NIAID/NIH. All mice were bred and maintained under pathogen-free conditions at an American Association for the Accreditation of Laboratory Animal Care accredited animal facility at the NIAID and housed in accordance with the procedures outlined in the Guide for the Care and Use of Laboratory Animals under an animal study proposal approved by the NIAID Animal Care and Use Committee. All mice used in experiments were male and used between 3–5 weeks of age so as to focus on acute muscle pathology.

Complete Blood Counts

Retro-orbital bleeding into EDTA-coated microvette tubes was performed on isoflurane-anesthetized mice. Leukocyte counts and differentials were determined using the HEMAVET® HV 950Fs Analyzer through the NCI Pathology and Hematology Laboratory, Frederick, MD.

Preparation of muscle tissue for histology and pathology scoring.

Mice were anaesthetized with isoflurane inhalation anesthesia and euthanized by cervical dislocation prior to dissection of hind limb skeletal muscles [23]. Quadriceps muscle tissues were processed as per a standard protocol. Briefly, the tissues were fixed in 10% neutral buffered formalin, sectioned, and blocked in paraffin for histological analysis. Tissue sections (5 μm) were stained with hematoxylin and eosin (H&E) for routine histopathology and then evaluated by a board-certified veterinary pathologist in a blinded manner. Sections were evaluated on six separate criteria: inflammation, myofiber degeneration-necrosis, myofiber regeneration, fibrosis, edema, mineralization. Each criterion was evaluated on a scale of 0 (absent/within normal limits), 1 (< 10% of tissue section affected), 2 (10–50% of tissue affected), or 3 (> 50% of tissue affected). All slides were examined using an Olympus BX51 light microscope and photomicrographs were taken using an Olympus DP73 camera.

Immunohistochemistry

Formalin-fixed paraffin-embedded quadricep muscle tissue sections were used to perform immunohistochemical staining using a polyclonal antibody, goat anti-eosinophil major basic protein (Santa Cruz, TX, sc-33938) at a dilution of 1:500. Staining was carried out on a Leica Bond Rx Automated Stainer platform (Leica Biosystems, Inc., Buffalo Grove, IL) according to manufacturer-supplied protocols. Briefly, 5 μm-thick sections were baked, deparaffinized and rehydrated. Heat-induced epitope retrieval (HIER) was performed using Leica Epitope Retrieval Solution 1, pH 6.0, heated to 100°C for 20 minutes. The slides were then incubated with hydrogen peroxide to quench endogenous peroxidase activity prior to applying the primary antibody followed by a biotinylated rabbit anti-goat secondary antibody (Abcam, Cambridge, MA, ab6740) at a dilution of 1:100. DAB chromogen detection was completed using the Bond Polymer Refine Detection kit and counter-stained using hematoxylin (Leica Biosystems, Inc., Buffalo Grove, IL). Slides were then cleared through gradient alcohol and xylene washes prior to mounting and coverslipping. Sections were examined by a board-certified veterinary pathologist using an Olympus BX51 light microscope and photomicrographs were taken using an Olympus DP73 camera.

Identification and characterization of eosinophils by flow cytometry and fluorescent activated cell sorting (FACS).

Mice were anaesthetized with isoflurane inhalation anesthesia and euthanized by cervical dislocation prior to dissection of hind limb skeletal muscles [23]. Muscle single cell suspensions were generated as previously described [23]. Briefly, skeletal muscle tissues were digested with type II collagenase and dispase. Cells were further separated by repeated passage through a 20G needle and filtered through a 40-micron cell strainer. Muscle single cell suspensions were frozen at 10 million cells per mL of freezing media (10% dimethyl sulfoxide in 90% Fetal Bovine Serum) at −80°C prior to analysis by flow cytometry. To quantify levels of eosinophils in the bone marrow, single-cell suspensions were prepared as previously described [24]. Briefly, bone marrow cells were isolated by flushing mouse femurs and tibiae with PBS. Red blood cells were eliminated by hypotonic lysis. Prior to analysis by flow cytometry, bone marrow single cell suspensions were frozen at −80°C in freezing media as described above. For analysis by flow cytometry, the following fluorochrome-conjugated antibodies and dyes were used: Live-Dead Fixable Aqua, MHC-II (Thermo Fisher); Gr1, Siglec-F, CD125 (BD Biosciences); CD45, CD11c, and CCR3 (Biolegend). CD16/32 (BD Biosciences) was used to block non-specific Fc Binding. A minimum of 200,000 events were collected on an LSR-II Flow Cytometer (BD Biosciences) and data were analyzed using Flow Jo (Tree Star). Eosinophils in muscle tissue were identified as LiveCD45⁺CD11c⁻Gr1⁻Siglec-F⁺MHC-II^low cells and verified by modified Giemsa staining of cytospin preparations (Diff-Quik, Thermo Scientific). Specific populations were identified by comparing fluorophore signals in samples with those of fluorescent minus one (FMO) and isotype antibody controls.

Serum and Muscle Tissue Cytokine Measurements

Whole blood was collected from isoflurane-anesthetized mice by the retro-orbital route. Blood allowed to clot for up to 30 minutes at room temperature. Samples were centrifuged at 3,000 rpm for 10 minutes and serum was stored at −80°C. Cytokines were measured by ELISA following manufacturer’s instructions. Quadriceps muscle tissues were removed from euthanized mice that had been completely exsanguinated. Quadriceps muscle tissue was flash frozen in liquid nitrogen-chilled isopentane and stored at −80°C. Subsequently, muscle tissue was homogenized in lysis buffer (100 mM Tris pH 7.4, 150 mM NaCl, 1mM EGTA, 1mM EDTA, 1% Triton X-100, 0.5% sodium deoxycholate, 1 mM phenyl-methyl sulfonyl fluoride and 1 tablet of protease inhibitor cocktail (Roche #04693159001)) per 10 mL of lysis buffer. Cytokine concentrations (pg / mg muscle protein) were measured by ELISA (R&D System) in 80 μg of protein, as determined by BCA assay (Thermo Fisher Scientific).

Serum Creatine Kinase Assay.

Sera levels of creatine kinase (CK) activity were analyzed by multiple-rate testing using Vitros® Creatine Kinase slides on the Vitros® 250 Chemistry Analyzer. To measure creatine kinase activity, each slide containing the creatine kinase substrate was loaded with 11 microliters of serum and incubated for 5 minutes. During incubation, reflection densities are recorded in a multiple-point rate. The rate of change in reflection density was monitored and the rate of change in reflection was then converted to enzymatic activity. For presentation, all values were normalized to the median value determined for wild-type sera samples, which was set at 1.0.

Muscle Fiber Size, location of nuclei, and detection of Pax7⁺ Stem Cells.

Cross-sections of frozen quadriceps muscle tissue were prepared as described [25] and stained with rabbit polyclonal anti-laminin (1:200, AbCam Cat No. ab11575), anti-Pax7 (1:75, Developmental Study Hybridoma Bank Cat No. Pax7-c), donkey anti-mouse IgG-Alexa 594 conjugate (Thermo Fisher Cat No. R37115), goat anti-rabbit IgG-Alexa488 conjugate (Thermo Fisher Cat No. R37116), and DAPI to visualize the nuclei. Imaging of the muscle tissue was performed using a Leica DMI6000 widefield microscope equipped with a 20×/0.7NA objective, a cooled DFC350FX monochrome camera, appropriate filter cubes, and an EL6000 light source. Tiled mosaic images were taken of whole tissue and then merged using the Navigator feature in the LAS × (Leica) software. Post-processing analysis was performed using Imaris software (Bitplane). To calculate the myofiber area and relative localization of myofiber nuclei, the “cell” module was used to first demarcate individual fibers which utilized the anti-laminin stain to delineate the membrane of each fiber followed by filtering by fiber area as well as degree of oblateness. Nuclei were identified and the shortest distance between a single nucleus and the lamnin-stained membrane was assessed for each fiber. If more than one nucleus was detected per fiber, the mean was calculated. Centrally-nucleated myofibers were determined to be those with nuclear staining at an average distance of 3 μm or greater from the anti-laminin-stained membrane. Pax7⁺ satellite cells were quantified per myofiber.

Statistical analysis.

Statistical significance was evaluated by ANOVA with multiple test corrections (GraphPad PRISM 7.0).

Results

Tissue pathology and quantitative evaluation of eosinophils in quadriceps muscle from wild-type, mdx, and eosinophil-deficient PHIL and mdx.PHIL mice.

Eosinophils were detected in muscle tissue from human subjects with DMD [19, 20] and in quadriceps and diaphragm muscles from mdx mice that phenocopy this disease [17–19]. However, the role of eosinophils in driving muscle damage has not been fully elucidated. Here we examine histological sections of quadriceps muscles from 4-week-old male wild-type, mdx, and mdx.PHIL mice; the last of this group are mdx mice that carry a transgene that promotes eosinophil ablation during hematopoiesis in vivo [21]. In contrast to what we observe in quadriceps muscle tissue from wild-type mice [Fig. 1A], the cross-section from the mdx mouse features abundant focal lesions [Fig. 1B, at arrowheads]. In addition, muscle tissue from mdx mice exhibited prominent regions of tissue edema, acute myofiber degeneration, necrosis, cytolysis, and leukocyte infiltration, with some evidence of myofiber regeneration, i.e., myofibers with central nuclei [Fig 1C, region enlarged from filled arrowhead in Fig 1B; also Suppl. Figs 1A–1C]. As shown in Fig. 1D, eosinophils both as single cells and in clusters are integral components of muscle lesions, as revealed by immunohistochemical localization with anti-eosinophil major basic protein-1 (anti-MBP-1). Muscle lesions were also prominent in cross-sections of quadriceps muscle tissue from mdx.PHIL mice, which harbor the PHIL transgene that ablates the eosinophil lineage [Fig. 1E]. The lesion at the filled arrowhead is enlarged in Fig. 1F, which features myofiber necrosis and leukocyte infiltration. However, as shown in Fig. 1G, immuno-histochemical staining with anti-MBP-1 reveals no eosinophils either in or around the muscle lesion.

Fig. 1. Histopathology and quantitative determination of eosinophils in quadriceps muscle from wild-type, mdx, eosinophil-deficient PHIL and mdx.PHIL mice.

A. Hematoxylin and eosin (H&E)-stained cross-sections of quadriceps muscles from 4-week-old male wild-type and B. 4-week-old mdx mice; in B., multiple regions of pathology as indicated by arrowheads, original magnifications, 4×. C. Enlargement of muscle lesion shown at filled arrowhead in B. featuring pronounced edema, acute myofiber necrosis, vacuolated cytoplasm, and leukocyte infiltration; original magnification, 20×; see also Suppl. Fig. 1A–1C. D. Immunohistochemical staining with anti-eosinophil major basic protein-1 (anti-MBP-1) documenting eosinophil infiltration. E. H&E-stained cross-section of quadriceps muscle tissue from 4-week-old male mdx.PHIL mouse, original magnification, 4×. F. Enlargement of muscle lesion shown at filled arrowhead in E. featuring acute necrosis and regenerating myofibers; original magnification, 20×. G. Immunohistochemical staining as in D.; no eosinophils detected in association with muscle lesion. H. Eosinophils (CD45⁺CD11c⁻Gr1⁻MHCII⁻SiglecF⁺ cells) isolated from quadriceps muscle by fluorescence-activated cell sorting (FACS) and stained with modified Giemsa. I. Eosinophils per mm² determined by visual inspection of anti-MBP-1 stained slides; 4–9 mice per genotype, ***p < 0.001, 1-way ANOVA. J. Eosinophils as % of live cells determined by flow cytometry (see Fig. 1H and Suppl. Fig. 1d), n = 2–10 mice per genotype, ***p < 0.001 one-way ANOVA.

Quantitative evaluation of eosinophils in muscle tissue was performed by visual inspection of anti-MBP-1-stained sections and by flow cytometry performed on single cell suspensions prepared from muscle tissue; eosinophils were identified as CD45⁺CD11c⁻MHCII^loGr1⁻SiglecF⁺ cells [Fig. 1H., and Suppl. Fig. 1D.]. Few eosinophils (2.3 ± 1.3 / mm²) were detected in muscle tissue from wild-type mice, and none detected in PHIL or mdx.PHIL mice, while 29 ± 18 eosinophils / mm² were detected in sections from mdx mice [***p < 0.001, Fig. 1I]. These findings were consistent with those determined by flow cytometry, in which eosinophils represented 1.20 ± 0.34% of the live cells from mdx mice, but < 0.1% of live cells detected in muscle from wild-type, eosinophil-deficient PHIL, or mdx.PHIL mice, and, respectively (***p < 0.001 of all vs. mdx, Fig. 1J).

From these findings we conclude that eosinophils are not the unique driving force behind acute muscle lesions, as lesions are prominent in quadriceps muscle of mdx.PHIL mice that are genetically devoid of eosinophilic leukocytes.

Tissue pathology and quantitative evaluation of eosinophils in quadriceps muscle from hyper-eosinophilic interleukin-5 (IL5) transgenic (tg) and mdx.IL5tg mice.

While eosinophil infiltration into the muscle tissue is a critical feature of DMD and the mdx mouse strain, it was not clear whether eosinophil infiltration alone, in dystrophin-sufficient mice, might also promote muscle damage. To address this question, we examined histological features in quadriceps muscle tissues from 4-week-old male IL5tg and mdx.IL5tg mice, which are mdx mice that carry a transgene that promotes eosinophilia and eosinophil activation in vivo [22]. While we detected no muscle lesions in the quadriceps sections from the IL5tg mouse [Fig. 2A], MBP-1⁺ eosinophils were detected among and between healthy myofibers throughout the muscle tissue [Fig. 2B and 2C]. By contrast, focal lesions were prominent throughout in sections of quadriceps muscle from mdx.IL5tg mice [Fig. 2D]; the enlargement in Fig. 2E features prominent leukocyte infiltration in association with multifocal mineralization of degenerate myofibers (at arrowheads). As shown in Fig. 2F, profound eosinophil infiltration was detected throughout, here in a region of pronounced myofiber degeneration.

Fig. 2. Histopathology and quantitative determination of eosinophils in quadriceps muscle from interleukin-5 (IL5) transgenic and mdx.IL5 transgenic mice.

A. H&E-stained cross-section of quadriceps muscle from a 4-week-old male IL5tg mouse; no lesions were detected, original magnification, 4×. B. Immunohistochemical staining with anti-MBP-1 and C. enlargement of area at arrowhead in B. revealing MBP-1⁺ eosinophils among healthy myofibers. D. H&E-stained cross section of quadriceps muscle from a 4-week-old male mdx.IL5tg mouse with lesions as indicated at arrowheads, original magnification, 4×. E. Lesion featuring extensive leukocyte infiltration in a region with mineralized myofibers at arrowheads; original magnification, 20×. F. Immunohistochemical staining as in B. and C. revealing numerous MBP-1⁺ eosinophils at muscle lesion. G. Eosinophils per mm² and H. Eosinophils as percent live cells were determined by visual inspection and flow cytometry, respectively as described in the legend to Fig. 1; n = 4–9 mice per group, ** p < 0.01, ***p < 0.001, Mann-Whitney U-test.

Quantitative evaluation of eosinophils in the muscle tissue was performed as described in Fig. 1. Eosinophils detected by visual inspection included 108 ± 16 / mm² in IL5tg mice and 337 ± 116 / mm² in mdx.IL5tg mice (**p < 0.01); of note, this represents 3.7-fold and 12-fold more eosinophils per unit area than detected in the sections from mdx mice, respectively (see Fig. 1I). Likewise, eosinophils represented 4.0 ± 1.1% and 20.0 ± 7.6% of the live cells in muscle tissue of IL5tg and mdx.IL5tg mice, respectively. These values, represent 3.3-fold and 17-fold more eosinophils in IL5tg and mdx.IL5tg mice, respectively, than were detected in muscle tissue from mdx mice (see Fig. 1J), and are consistent with the findings from immunohistochemical staining and visual inspection.

From these findings, we can conclude that eosinophils alone, even in large numbers, do not a priori generate muscle lesions in the absence of additional signals such as those provided (directly or indirectly) by dystrophin deficiency. The relationship between eosinophil infiltration and the extent of muscle lesions from all strains will be examined quantitatively in the section to follow.

Quantitative indicators of muscle pathology.

We have shown that muscle lesions are prominent in mdx mice that are devoid of eosinophils (i.e., mdx.PHIL). Likewise, we find that dystrophin-sufficient mice with eosinophils infiltrating muscle tissue do not develop these lesions (i.e., IL5tg). As such, it will be critical to evaluate these findings quantitatively to define fully the association (or lack thereof) between muscle lesions and eosinophil infiltration. As a first step, the histological sections prepared from quadriceps muscle tissue from 4-week-old male mice from all six genotypes were evaluated quantitatively with a pathological scoring system. Three to six sections per genotype were coded and scored by a skilled investigator who evaluated them in “blinded” fashion. The scoring system included six parameters: inflammation, edema, myofiber degeneration, fibrosis, myofiber mineralization, and myofiber regeneration. Each section was scored on a 0–3 scale of least to most tissue involvement (see Methods) for a maximum possible score of 18 points. As shown in Fig. 3A, muscle tissue sections from wild-type and PHIL mice displayed no pathology (score 0) and those from IL5tg mice scored 1.0 ± 0.8 points due to the inflammation sub-score and eosinophil influx as noted above (Fig. 2B). By contrast, mdx and all other mdx-derived dystrophin-deficient strains displayed elevated pathology scores. The score for eosinophil-deficient mdx.PHIL mice (6.3 ± 0.6 points) was somewhat lower than that for parent mdx mice (9.7 ± 0.8 points); however, no significant difference was noted between scores determined for eosinophil-deficient mdx.PHIL and eosinophil-enriched mdx.IL5tg (7.8 ± 1.0 points).

Fig. 3. Quantitative indicators of muscle pathology.

A. Muscle histopathology score. H&E-stained sections from quadriceps muscles from 4-week-old male mice were scored on individual criteria including inflammation, edema, necrosis-degeneration, mineralization, myofiber regeneration, and fibrosis, each on a 0–3 intensity scale with a maximum potential score of 18 overall, see Methods for additional details; n = 3–6 mice per genotype, **p < 0.01, ***p < 0.001 1-way ANOVA. B. Myofiber area: percent of myofibers from wild-type (wt) vs. mdx quadriceps muscle identified within the size ranges as indicated; **p < 0.01, *p < 0.05, 2-way ANOVA, n = 1.2 × 10⁴ myofibers (wild-type), 0.7 × 10⁴ myofibers (mdx), 2–3 mice per genotype. C. Myofiber area: percent of myofibers from eosinophil-deficient PHIL vs. mdx.PHIL quadriceps muscle identified within the size ranges as indicated; ***p < 0.001, *p < 0.05, 2-way ANOVA, n = 1.1 × 10⁴ myofibers (PHIL), 0.7 × 10⁴ myofibers (mdx.PHIL), 3 mice per genotype. D. Central nuclei: percent of myofibers from wild-type (wt), mdx, PHIL, and mdx.PHIL quadriceps muscle with centrally-placed nuclei; ***p < 0.001, 1-way ANOVA, 2–3 mice per genotype. E. Serum creatine kinase: Values shown were normalized to a single value from wild-type (wt); n = 15–20 mice per genotype, *p < 0.05, ***p < 0.001, ns, no significant difference.

Distribution of myofiber size has been used as a measure to evaluate healthy vs. degenerating and regenerating muscle tissue [26]. As shown in Fig. 3B, areas determined for myofibers from quadriceps muscle from wild-type mice (n = 3 mice, 1.2 × 10⁴ myofibers evaluated, binned in groups of 1000) generate a normal distribution; the mean myofiber area calculates to 2815 ± 1540 μm²). By contrast, the myofibers evaluated in muscle tissue from mdx mice (n = 2 mice, 0.7 × 10⁴ myofibers evaluated) are not normally distributed and display prominent skewing toward both lower and higher size ranges as anticipated. A similar set of curves was generated to provide a comparison between fiber sizes from muscles of eosinophil-deficient PHIL and mdx.PHIL mice [Fig. 3C]. The myofiber sizes for PHIL mice (n = 3 mice, 1.1 × 10⁴ myofibers evaluated, binned as above) generate a normal distribution, while those from the dystrophin-deficient mdx.PHIL mice (n = 3 mice, 0.7 × 10⁴ myofibers) generate a curve similar to that shown in Fig. 3B for the mdx mice, with size heterogeneity and skewing toward both the lower and higher ends of the size ranges.

The nucleus of a healthy mature myofiber is eccentrically placed and located adjacent to the cell membrane; regenerating myofibers can be identified by nuclei located internally, or centrally within the myofiber [27]. Distances between nuclei and cell membrane were evaluated in quadriceps muscle from wild-type, mdx, PHIL and mdx-PHIL mice (n = 2–3 mice per group, 200–8000 myofibers per muscle section). As shown in Fig. 3D, most (70–95%) of the myofibers in all four strains had nuclei located adjacent or nearly adjacent to the membrane. A larger proportion of the myofibers from the mdx (27 ± 2.4%) and mdx.PHIL (30 ± 1.9%) had centrally-placed nuclei indicating muscle regeneration; only 5.8 ± 0.9% of the myofibers from the wild-type and 4.7 ± 1.0 % from the PHIL mice were placed within this group (***p < 0.001; see also [Suppl. Fig. 2A and 2B]).

We performed a similar analysis that included immunohistochemical staining with anti-PAX7 to detect muscle satellite cells. Although PAX7⁺ cells were detected in quadriceps muscle from all four strains, no significant differences were detected (Suppl. Fig. 2C and 2D.).

Finally, serum levels of the myofiber-derived enzyme, creatine kinase (CK), were evaluated as an indirect measure of acute muscle damage [28]. Consistent with our earlier findings, the presence or absence of eosinophils had no impact on serum CK measurements. As shown in Fig. 3E, serum CK levels in mdx mice were more than 2-fold greater than those in wild-type mice (2.23 ± 0.62 vs. 1.00 ± 0.48, normalized units), as were levels in mdx.PHIL vs. parent PHIL mice (***p < 0.001 for each pair). Interestingly, serum CK measured for mdx.IL5tg mice was comparatively low, although peripheral blood eosinophilia and elevated levels of cationic proteins in serum may bind to and interfere with the components and readout of this assay.

Taken together, these measures of muscle pathology confirm that while eosinophils are present in tissue lesions associated with dystrophin deficiency, they are not a driving force promoting tissue destruction.

Mechanisms of eosinophil recruitment to muscle tissue of mdx mice.

Eosinophil recruitment to and infiltration into peripheral tissues has been examined primarily in disorders involving Th2 cytokines, notably allergy and helminth infection [29]. These conditions are typically associated with elevated levels of the eosinophil-active cytokine, IL5, which induces expansion of eosinophil progenitors in bone marrow and promotes eosinophil survival in the periphery. Interestingly, while the IL5 transgenic crossed strains are characterized by eosinophilia in both peripheral blood [Fig. 4A] and bone marrow [Fig. 4B], eosinophils in these compartments in 4-week-old male mdx mice are at levels that are indistinguishable from wild-type controls. Likewise, serum IL5 levels were elevated only in the transgenic strains and not in mdx or mdx.PHIL mice [Fig. 4C]; likewise, no IL-4 and no IL-13 was detected above baseline levels (data not shown), although this was not the case for muscle tissue levels of IL-4, addressed in the section to follow.

Fig. 4. Eosinophil recruitment to muscle tissue.

A. Peripheral blood and B. bone marrow eosinophils in mdx mice are indistinguishable from the wild-type; elevated levels are observed in IL5tg mice (n = 2–9 mice per genotype, ***p < 0.001). C. Serum IL-5 in wild-type (wt), mdx, PHIL, mdx.PHIL, IL5tg and mdx.IL5tg mice; n = 8–10 mice per genotype, ***p < 0.001. D. Eotaxin-1/CCL11 was detected in muscle tissue from mdx, mdx.PHIL and mdx.IL5tg mice at levels that exceed those detected in wt, PHIL, and IL5tg mice, respectively; n = 5–9 mice per genotype, **p < 0.01 (ANOVA). E. Eotaxin-1 in muscle tissue correlates inversely with age specifically in the mdx strain; R² = 0.72, ***p < 0.0001. F. Serum eotaxin-1 in wild-type (wt), mdx, PHIL, mdx.PHIL, IL5tg and mdx.IL5tg mice; n = 8–9 mice per genotype, ***p < 0.001. G. RANTES / CCL5 is detected in muscle tissue from mdx, mdx.PHIL and mdx.IL5tg mice at levels that exceed those detected in wt, PHIL, and IL5tg mice, respectively; n = 5–9 mice per genotype, *p < 0.05, **p < 0.01, ***p < 0.001; 2-way ANOVA. H. Muscle eosinophils express CCR3. I. MFI for CCR3 is higher on eosinophils from the mdx-derived strains (mdx and mdx.IL5tg) than on those from the IL5tg alone (**p < 0.01; n = 8–10 mice per genotype).

To understand how eosinophils may be recruited to muscle tissue, we explored tissue levels of critical eosinophil chemoattractants. As shown in Fig. 4D, the three mdx-derived strains (mdx, mdx.PHIL and mdx.IL5tg) displayed elevated concentrations of eotaxin-1 (CCL11; pg/mg muscle lysate) at 4 weeks of age compared to their respective dystrophin-sufficient counterparts. Interestingly, the concentration of eotaxin-1 in muscle lysates from mdx mice correlated inversely with the age of the mouse [Fig. 4E]. These results suggest that eosinophil activation and recruitment may be initiated very early during the mouse lifespan. Inbred mice are mobile as early as 6 days after birth, and they may already be damaging skeletal muscle; interestingly, early eosinophil recruitment secondary to a “pre-dystrophic” state, has been suggested for Limb-girdle muscular dystrophy 2A / Calpain-3 deficiency [30], another form of muscular dystrophy ([31, 32]; see Discussion. We note that serum levels of eotaxin-1 do not correlate with those detected in muscle tissue [Fig. 4F]. The eosinophil chemoattractant RANTES (CCL5) was also detected in lysates from quadriceps muscle of mdx and mdx-derived mouse strains [Fig. 4G] but not in lysates from their dystrophin-sufficient counterparts. Furthermore, we identified CCR3, the receptor for eotaxin-1 and RANTES [33] on eosinophils in muscle tissue from mdx, mdx.IL5tg and IL5tg strains [Fig. 4H]. While the we detected no differences in percent CCR3-positive cells, MFIs differed significantly; eosinophils from both mdx-derived strains displayed higher MFIs for CCR3 than did eosinophils from the dystrophin-functional, IL5tg mice (mdx vs. IL5tg; mdx.IL5tg vs. IL5tg, **p < 0.01) [Fig. 4I].

Eosinophils contribute immunomodulatory mediators to the muscle microenvinronment.

Recent explorations of eosinophil function focus on their role in promoting homeostasis via release of immunomodulatory mediators into the local microenvironment [34, 35]. Given the number of studies that have revealed a role of IL-4 as a prominent mediator of eosinophil function in health and disease (for example, [36–38]), we examined muscle tissue from all six strains for levels of this cytokine. Of interest, we did detect relatively small (~20–25%) but statistically significant increases in IL-4 concentrations in muscle lysates with prominent eosinophil infiltration, ie., quadriceps muscle from mdx, IL5tg and mdx.IL5tg mice (*p < 0.05, **p < 0.01 vs. wt, PHIL, and mdx.PHIL strains; Fig. 5A). However, we note that increases in immunoreactive IL-4 were not proportional to absolute eosinophil concentration in muscle tissue. Furthermore, no increases in IL-13 over background were detected [Fig. 5B]. Interestingly, we did detect very high concentrations of IL-1Ra/IL-1F3 in muscle tissue from the highly hyper-eosinophilic mdx.IL5tg mice (3513 ± 1138 pg/mg muscle tissue; Fig. 5C). IL-1Ra is an anti-inflammatory mediator and natural inhibitor of IL-1β; Suguwara and colleagues [39] recently reported that eosinophils in the gastrointestinal tract secreted large quantities of IL-1Ra and noted that this factor was critical for suppressing Th17 cells and for maintaining gut homeostasis.

Figure 5. Cytokines detected in muscle tissue lysates.

Immunoreactive A. IL-4, B. IL-13 and C. IL-1RA/IL-1F3 detected in muscle tissue from wild-type (wt), mdx, PHIL, mdx.PHIL, IL5tg and mdx.IL5tg mice; n = 5–9 mice per genotype, *p < 0.05, **p < 0.01, ***p < 0.005, 1-way ANOVA.

Discussion

In this study, we utilized the mdx mouse model of Duchenne Muscular Dystrophy (DMD) to examine eosinophilic inflammation and its association with acute muscle damage. To do so, we generated mdx mice devoid of eosinophils (mdx.PHIL) and mdx with hyper-eosinophilia (mdx.IL5tg) and evaluated four critical measures of muscle damage, focusing on quadriceps lesions in 4 week-old male mice: (1) acute histopathology, (2) size distribution of myofibers, (3) fraction of regenerating myofibers, and (4) serum creatine kinase. From these findings, we concluded that, while eosinophils are prominent in these muscle lesions, they do not drive the acute muscle damage. Specifically, we found that genetic ablation of the eosinophil lineage in mdx.PHIL mice had no consistent impact on quantitative measures of muscle damage when compared directly to parent mdx mice. Moreover, our studies with IL5Tg and mdx.IL5tg mice demonstrate that eosinophil infiltration does not a priori promote muscle destruction; likewise, the degree of eosinophil infiltration detected within quadriceps lesions of dystrophin-deficient mice does not correlate with the extent of tissue damage.

These findings challenge the long-held perception of eosinophils as uniformly cytotoxic [29, 35]. Eosinophilic infiltration into the muscle tissue has been described in association with an extensive range of clinical pathologies, including acute myocarditis, myositis with and without infection, distinct myalgias (e.g. Toxic Oil and Eosinophil-Myalgia syndromes), as well as the muscular dystrophies [40–44], including Limb Girdle Muscular Dystrophy Type 2A (LGMD2A) / Calpain-3 (CLPN-3) deficiency [30–32]. The therapeutic efficacy of corticosteroids and their impact on eosinophils in tissues has naturally led to the assumption that these cells are critical mediators of acute damage and dysfunction, although this has not been formally proven in any of these disorders. Interestingly, eosinophil infiltration in the muscle tissue without (or prior to) histopathological changes was reported Krahn and colleagues [30] in a series featuring six children with CLPN-3 deficiency. This has led to speculation that eosinophils may be recruited during a ‘pre-dystrophic’ stage of this disorder, when subtle myofiber defects generating signals may promote eosinophil accumulation even before overt histopathology can be detected. Similarly, pre-dystrophic myofiber defects may play a role in promoting eosinophil infiltration in DMD and in the mdx mouse model, even though no overt histopathology is in evidence.

The eosinophils in muscle lesions characteristic of DMD and the mdx mouse strain have also been the subject of significant exploration [17–19]. Interestingly, while eosinophils and the primary eosinophil secretory mediator, major basic protein-1 (MBP-1) both promote muscle cell lysis in vitro [19] their cytotoxic potential in vivo is somewhat less clear-cut. Wehling-Hendricks and colleagues [19] found that repeated administration of anti-CCR3 resulted in significant eosinophil depletion in association with diminished severity of associated muscle lesions; however, this depletion strategy may also have an impact on CCR3⁺ T lymphocytes and activated CCR3⁺ B-cells [45–48]. Likewise, genetic ablation of MBP-1, via generation of mdx.MBP-1^−/− mice, had limited impact on muscle damage in vivo [19]. New mouse models, such as PHIL [21] permit us to address issues focused on eosinophils and their role in promoting muscle pathology with greater precision.

Several groups have examined interactions between eosinophils and acute muscle damage using other experimental models. For example, Heredia and colleagues [49] generated acute damage in the mouse tibialis anterior muscle via injection of cobra cardiotoxin, a 6.8 kDa polypeptide that depolarizes the cell membrane and promotes contracture of skeletal muscle; in this model, acute injury was accompanied by myofiber regeneration observed within 8 days after injection. Among their conclusions, the authors found that eosinophils promoted muscle regeneration primarily via the release of pre-formed IL-4. These findings are consistent with the work of Horsley and colleagues [50] who first reported that IL-4 promoted skeletal muscle growth, and likewise with the “LIAR hypothesis” [35], which posits that eosinophils are not primarily cytotoxic but instead recruited in response to tissue damage to modulate local immunity and repair. Here, we identified small but statistically significant increases in immunoreactive IL-4 in all muscle tissues that include eosinophils over baseline (mdx, IL5tg, mdx.IL5tg), although IL-4 levels did not correlate with the extent of eosinophil infiltration nor was it associated specifically with dystrophin-deficiency (Fig. 5A). Furthermore, with respect to muscle regeneration, we note that our current findings revealed no differences absolute numbers of regenerating myofibers in mdx eosinophil-sufficient vs. mdx.PHIL eosinophil-deficient mice at the 4 week time point (Fig. 3). These findings require further evaluation, notably at later time points (i.e., 7–8 weeks) that feature more prominent evidence of muscle tissue regeneration and repair [51].

By contrast, it is interesting to note the findings of Diny and colleagues [37] who report that eosinophils promoted cardiac dilation and associated muscle dysfunction in a model of autoimmune myocarditis. The differences in the two disease models, distinct microenvironments, as well as the nature of eosinophil activation and recruitment, all require further consideration.

Finally, we found no peripheral eosinophilia and no elevated levels of serum IL-5 in 4-week-old mdx mice. This is notable, as eosinophil infiltration into tissues is frequently associated with Th2 lymphocyte activation and elevated levels of serum IL-5, followed by expansion of the eosinophil population in bone marrow and peripheral blood [29]. While peripheral eosinophilia has been reported in association with eosinophil-associated tissue myopathies [44], Shröder and colleagues [20] reported that that peripheral blood eosinophil counts often do not correspond directly with the degree of eosinophil infiltration in the muscle tissue. However, we recognize that while mouse and human eosinophils maintain many of the same major proteins and receptors and are functionally and structurally analogous, they are not fully identical [29, 52]; as such, the precise role of eosinophils in human disease may be somewhat different than that revealed here for dystrophin-deficient mouse strains.

We did detect elevated levels of eosinophil chemo-attractants eotaxin-1 (CCL11) and RANTES (CCL5) in the muscle tissues of all three dystrophin-deficient mouse strains (mdx, mdx.PHIL and mdx.IL5tg). Interestingly, levels of eotaxin-1 in muscle tissue of mdx mice correlate negatively with age; higher levels were detected among the 3-week-old mice, returning to near wild-type levels in 5-week-old mice. These findings suggest that eosinophils may be infiltrating muscle tissue at an early point in post-natal development, analogous what has been hypothesized for human calpain-3 deficiency as noted above [30]. Likewise, although we cannot detect IL-5 in serum or tissue of 4-week-old mice, we cannot rule out the possibility that IL5 contributes to eosinophil recruitment, perhaps also during a brief, time-limited period earlier in the mouse lifespan. We note that IL5-deficient (IL5^−/−) mice maintain homeostatic levels of eosinophils in the peripheral blood and bone marrow that are virtually indistinguishable from the wild-type [53], yet no eosinophils were detected in dystrophic muscle lesions in mdx.IL5^−/− mice (Sek, Rosenberg; unpublished findings).

In summary, we have determined that, although eosinophils are prominent in muscle infiltrates, these cells do not promote acute muscle damage in the mdx mouse model of Duchenne Muscular Dystrophy. We determined that manipulating the level of eosinophils in the muscle tissue either by systemic ablation or cytokine-mediated augmentation, had no major impact on the degree of acute muscle damage. As eosinophils have profound immunomodulatory capacity, ongoing studies will address the unique properties of eosinophils recruited to muscle tissue and their roles in promoting regeneration and muscle growth and tissue repair in vivo.

Key Points.

• Eosinophils are prominent in quadriceps lesions of dystrophin-deficient mdx mice.

• Muscle damage was evaluated quantitatively in mdx, mdx.PHIL, and mdx.IL5tg mice.

• In these models, eosinophil infiltration is not driving acute muscle pathology.