Inflammation in a severe model of dystrophic cardiomyopathy contains a high proportion of T cells that contribute to onset of pathology

Abstract

Duchenne muscular dystrophy (DMD) is a severe neuromuscular disorder with progressive muscle degeneration and cardiomyopathy leading to heart failure. The inflammatory environment in dystrophic skeletal muscle is well-studied, but little is known about inflammation in DMD cardiomyopathy due to the lack of adequate animal models. We recently developed the Fiona/dko mouse model, deficient for both dystrophin and utrophin, but containing a skeletal muscle specific expressing utrophin transgene allowing progression of dystrophic cardiomyopathy. This Fiona/dko model is the first DMD cardiomyopathy model to reproducibly progress to reduced cardiac contractile function by 9 months. In this study, we compared immune cell composition between Fiona/dko mice and their milder littermates that develop cardiac pathology, but do not demonstrate whole heart dysfunction. Flow cytometry analysis revealed that T cells constitute a significant proportion of the immune cell population in dystrophic hearts, in contrast to the known predominantly myeloid signature in dystrophic skeletal muscles. T cell infiltration precedes development of cardiac fibrosis and dysfunction in Fiona/dko mice. RNA sequencing of whole hearts after cardiac dysfunction shows increased expression of 68 genes related to T cell signaling in Fiona/dko compared to their milder littermates. Furthermore, depletion of circulating CD3⁺ T cells with a neutralizing antibody ameliorates early pathology in Fiona/dko hearts. Together, these data suggest a role for T cells in initiation and persistence of dystrophic cardiomyopathy. These findings highlight the distinct inflammatory environment in the dystrophic heart and provide new insights into DMD cardiomyopathy, paving the way for the future development of targeted anti-inflammatory therapies.

New & Noteworthy

Heart failure has become the leading cause of death in Duchenne muscular dystrophy, a progressive degenerative disease of striated muscles. This study highlights an inflammatory environment in dystrophic heart with a high proportion of T-cells that is distinct from the predominant myeloid inflammation in dystrophic skeletal muscles. T-cell related signaling is associated with the severity of cardiomyopathy and T-cells contribute to dystrophic cardiomyopathy onset. This data will inform optimal patient treatments that target cardiac inflammation.

Introduction

Duchenne muscular dystrophy (DMD) is a severe and progressive neuromuscular disorder and is the most common muscular dystrophy affecting 1:5000 males. This condition is caused by lack of dystrophin protein due to mutations in the X-linked DMD gene. Lack of this important structural protein leads to significant striated muscle damage, degeneration and weakness, markedly impacting patients’ quality of life. There is a high prevalence of cardiomyopathy in DMD, with approximately 90% of 18-year-old male patients, and up to 50% of female carriers exhibiting cardiac complications (1–7). Heart failure is also now the leading cause of death in DMD (1, 2, 5, 6). This progressive cardiac involvement indicates a dire need for a deeper understanding of disease mechanisms that will enable development of more effective treatments.

DMD cardiomyopathy and treatment outcomes have been difficult to study preclinically. This challenge existed because the most commonly used genotypic animal model, mdx mice, which have been used to dissect much of the pathophysiology and treatment approaches for dystrophic skeletal muscles, do not recapitulate dystrophic cardiomyopathy and do not progress to reduced ejection fraction. Mdx mice also deficient for the dystrophin paralog utrophin (“dko” mice) show early cardiomyopathy pathology similar to DMD patients, but succumb to their skeletal muscle disease before cardiomyopathy progresses (8). To overcome this barrier, our lab developed a mouse model of isolated DMD cardiomyopathy that we have shown to recapitulate the human progression from cardiomyopathy to reduced strain to reduced ejection fraction indicative of heart failure (9). These mice, called Fiona/dko (Fiona/Dmd^mdx;utrn^–/–), have whole-body deficiency of dystrophin and utrophin (dko) but then overexpress a human utrophin transgene driven by ACTA1 promoter specifically in skeletal muscles (“Fiona” transgene) (9). Thus, skeletal muscles are rescued by the overexpression of utrophin allowing the dko cardiomyopathy to progress. This model has enabled us to begin studying the pathogenesis of dystrophic cardiomyopathy and its progression.

A critical component of DMD pathology is the chronic inflammation that occurs as muscle cells break down. Damaged cell membranes release damage-associated molecular patterns, cytokines and chemokines, triggering an immune response that has been well-established to play a role in the pathology of skeletal muscle (10–21). However, due to the previous lack of an adequate dystrophic cardiomyopathy model, little is known about the inflammatory environment of the heart. Currently, corticosteroids are the standard-of-care treatment for DMD, most likely acting through their general anti-inflammatory mechanism of action (22–25). However, their long-term use is associated with numerous adverse effects, including osteoporosis, behavioral issues and Cushing syndrome (26). Therefore, there is a pressing need for novel and more specific anti-inflammatory therapies.

Inflammation is emerging as a pivotal pathogenic mechanism in DMD skeletal muscle, involving a predominantly myeloid signature consisting of macrophages, monocytes, neutrophils, eosinophils, and natural killer cells (19, 20, 27). Our previous studies demonstrate that over 90% of immune cells in both quadriceps and diaphragm skeletal muscles from 2 dystrophic models consist of myeloid cells representative of the innate immune system (28–30). It has even been suggested that macrophages are responsible for most of the membrane lysis and damage in mdx mouse skeletal muscles, as opposed to auto-mechanical damage due to the lack of dystrophin causing membrane instability (10, 19, 25). However, macrophages and LY6C^hi monocytes also contribute to skeletal muscle regeneration, making them a complicated therapeutic target (31, 32). In addition, small percentages of CD4 and CD8 positive T cells are present in dystrophic skeletal muscles and are known to amplify immune responses through the release of cytokines and recruitment of other immune cells via chemokines, worsening fibrosis and necrosis (12, 14, 16, 17, 33).

The role of T cells in driving DMD cardiomyopathy progression to heart failure has been difficult to elucidate due to lack of animal models that exhibit heart dysfunction. However, two studies have shown that T cells are present in dystrophic heart and suggest they can modulate pathology (21, 34). When mdx mice were treated long-term with human IgG, a treatment used clinically for autoimmune disorders to disrupt antibody signaling, fractional shortening and ejection fraction were increased while T cell and macrophage infiltration into the heart was decreased (21). Recently, T cells were also shown to be present in hearts from exercised mdx mice, a model that slightly augments cardiomyopathy in this very mild model (34). An inhibitor of an important protein involved in T cell activation, PKCƟ, was shown to be beneficial in exercised mdx mice by reducing all immune cells as well as necrosis and fibrosis in the heart, although this model still does not progress into cardiac dysfunction (34). T cells are also known to contribute to numerous other cardiac pathologies including myocardial infarction and myocarditis (35–42).

To begin to uncover the contribution of inflammation to dystrophic cardiomyopathy in a model that progresses to heart failure, we investigated cardiac immune cells and gene expression in Fiona/dko mice compared to their less severe littermates, Het/Fiona (Fiona/Dmd^mdx;utrn^+/−) mice, which retain expression from one copy of utrophin in the heart and do not progress into reduced whole heart function (9). We observed that the heart immune cell population contains a high proportion of T-cells prior to dysfunction and that both myeloid and T-cells persist throughout cardiomyopathy progression and functional decline. This data suggests that inflammation in dystrophic heart is remarkably different from dystrophic skeletal muscles that contain predominantly myeloid cells with a peak at the onset of pathology followed by a dramatic decline during disease progression (28–30). We compared T cell signaling between the two models and targeted T cells in Fiona/dko mice to begin to uncover their contribution to the onset of dystrophic cardiomyopathy. These studies will inform future therapeutic designs targeting pro-inflammatory and dysregulated T cells in DMD cardiomyopathy and heart failure.

Materials and Methods:

Sex as a biological variable.

Male and female animals were used whenever possible in this study and findings were similar for both sexes.

Mice.

To generate Fiona/dko (Fiona/Dmd^mdx Utrn^–/–) experimental mice containing a full-length human utrophin cDNA under control of the Acta1 skeletal muscle-specific promoter (Tg (ACTA1-Utrn) 2^Ked) (“Fiona”) on a dystrophin/utrophin-deficient (dko) background, Fiona/Dmd^mdx Utrn^+/− (Het/Fiona) mice were mated with Dmd^mdx Utrn^+/− (Het) mice (9). The Fiona transgene was detected by PCR with 5′-GTCAGGAGGGGCAAACCCGC-3′ (Utr_TG_For) and 5′-GTCGCTGCCCTTCTCGAGCC-3′ (Utr_TG_Rev) primers. The utrophin knockout allele was detected with 5′-GACAAACTGTCAGTTCTTAAG-3′ (UTRF1) and 5′-ACGAGACTAGTGAGACGTGC-3′ (NeoR) and the wild-type utrophin allele was detected with 5′-GTGAAGGATGTCATGAAAG-3′ (PU65) and 5′-TGAAGTCCGAAAGAGATACC-3′ (Intron 7). Both mouse lines have been previously backcrossed for many generations over decades with C57BL/10-mdx mice.

Generation of single-cell suspensions from cardiac muscles.

Single-cell suspensions were generated from 3- and 9-month-old Het/Fiona and Fiona/dko hearts for flow cytometric analysis of immune cells. Briefly, hearts (n = 3-month-old Het/Fiona: 3M, 4F; 3-month-old Fiona/dko: 3M, 3F; 9-month-old Het/Fiona: 10M, 5F; 9-month-old Fiona/dko: 8M, 7F) were dissected, rinsed in cold DPBS, and finely minced with razor blades. The heart from each individual mouse was processed and analyzed as an independent sample. Following dissociation, muscles were digested with 10 mL/g of digestion buffer composed of DMEM (Thermo Fisher Scientific, 11995–065), 0.02% Collagenase P (Roche, 11213857001), and 0.1% RQ1 DNase (Promega, M6101) in a 37°C water bath for 30 minutes. After digestion, the resulting suspension was passed through a 70 μm filter then a 40 μm filter, which filters out cardiomyocytes. The cells from the filtrate were fixed in 1% paraformaldehyde on ice for 10 minutes and then rinsed in cold flow buffer and centrifuged. The supernatant was discarded and the cell pellet was resuspended in cold flow buffer. All single-cell suspensions were stored at 4°C following fixation for less than 2 weeks before staining for flow cytometry.

Flow cytometric analysis of cardiac immune cells.

Flow cytometry staining was done separately for the myeloid and lymphoid panels. Flow cytometry antibodies utilized for the myeloid cell panel staining were as follows: CD45 (PE-Cy7; Thermo Fisher Scientific, 25045182), CD11b (APC; BioLegend, 101212), LY6G (APC/FIRE750; BioLegend, 127652), LY6C (eFluor450; Thermo Fisher Scientific, 48593282), CD64 (BV605; BioLegend, 139323), CD206 (PerCP-eFluor710; Thermo Fisher Scientific, 46206182), CCR2 (FITC; BioLegend, 150608) and CCR5 (PE; BioLegend, 107006). Flow cytometry antibodies utilized for the lymphocyte panel staining were as follows: CD45 (PE-Cy7; Thermo Fisher Scientific, 25045182), CD3 (violetFluor 500; Tonbo, 85–0032-U100), CD4 (Super Bright 600; Invitrogen, 63–0040-82), CD19 (SparkBlue 550; BioLegend, 115566), and FoxP3 (APC; Invitrogen, 17–5773-82). The markers were used to identify leukocytes (CD45⁺), myeloid cells (CD45⁺CD11b⁺), T cells (CD45⁺CD3⁺), neutrophils (CD45⁺CD11b⁺LY6G⁺), macrophages/monocytes (CD45⁺CD11b⁺LY6G–CD64⁺), and T cell and macrophage subtypes in the single-cell suspensions. Corresponding isotype controls were included in all experiments. Cells were permeabilized prior to CD206 and FoxP3 staining using DPBS-solubilized 0.5% Tween-20 (Sigma-Aldrich, P1379). Experiments were performed using a Cytek Northern Lights Flow Cytometer, and data were analyzed with FlowJo software version 10.9.0 (Becton Dickinson).

Immune cell isolation from blood.

For blood leukocyte isolation, 50 μL of blood obtained via submandibular blood collection was incubated with 2 mL of RBC Lysis Buffer (Thermo Fisher; 00433357) for 5 minutes. Cells were then fixed with 1% paraformaldehyde on ice for 10 minutes, rinsed in cold flow buffer and centrifuged. The supernatant was discarded and the cell pellet was resuspended in cold flow buffer. All single-cell suspensions were kept at 4°C following fixation for less than 2 weeks before staining for flow cytometry.

Flow cytometric analysis of blood immune cells.

Flow cytometry antibodies utilized for staining blood immune cells were as follows: CD45 (Super Bright 600; Invitrogen, 63045180), CD3 (FITC; BioLegend, 100306), CD4 (PE-Cy7; BioLegend, 100528) and CD8 (BV650; BioLegend, 100742). The markers were used to identify leukocytes (CD45⁺), T cells (CD3⁺), helper T cells (CD3⁺CD4⁺CD8⁻) and cytotoxic T cells (CD3⁺CD8⁺CD4⁻). Corresponding isotype controls were included in all experiments. Experiments were performed using a Cytek Northern Lights Flow Cytometer, and data were analyzed with FlowJo software version 10.9.0 (Becton Dickinson).

Mouse treatments.

8-week-old Fiona/dko mice were injected intraperitoneally with either anti-CD3 antibody (BioXCell, BE0002) (n = 2M, 1F) or isotype antibody (BioXCell, BE0090) (n = 3M). The treatment regimen began with 3 successive days of 150 μg antibody in 50 μL sterile saline injections once per day followed by twice weekly injections of 200 μg antibody in 50 μL sterile saline for 4 weeks, resulting in a total treatment course of 5 weeks. Mice were housed up to 5 per cage and euthanized by cervical dislocation prior to dissection.

Histology and immunofluorescence and immunohistochemical analysis.

Hearts were isolated from 2-, 3-, 6-, 9-, and 12-month-old Fiona/dko mice (n = 2-month: 2M, 1F; 3-month: 1M, 2F; 6-month: 1M, 2F; 9-month: 1M, 2F), control Het and C57BL/10 mice and 3 month-old antibody-treated Fiona/dko mice (n = isotype: 3M; anti-CD3 antibody: 2M, 1F) and embedded in optimal cutting temperature (OCT) compound, frozen on liquid nitrogen-cooled isopentane, and cut into 8 μm sections on a cryostat (Bright Instruments). Quality control of sections was performed by hematoxylin and eosin (H&E) staining for overall histology. Immunofluorescence for damage and fibrosis was performed with 1:40 rabbit anti–mouse fibronectin primary antibody (Abcam, 23750) and 1:200 Alexa Fluor 555–conjugated goat anti-rabbit secondary antibody (Invitrogen, A-21429) together with 1:100 AlexaFluor 488-conjugated goat anti-mouse IgG (Invitrogen A-11029) to detect endogenous IgG infiltration within the myocardium. For lymphoid and myeloid immune cell immunohistochemistry localization, sections were incubated with 1:10 rabbit anti-mouse monoclonal CD3 antibody (Abcam, ab135372), 1:50 rat anti–mouse monoclonal CD11b antibody (BD Pharmingen, 550282) or 1:50 rat anti–mouse monoclonal CD45 antibody (BD Pharmingen, 550539); then incubated with either 1:200 HRP-conjugated goat anti-rabbit (Jackson ImmunoResearch, 111–035-144) or 1:200 HRP-conjugated goat anti-rat (Invitrogen, A10549) secondary antibodies, as appropriate; then incubated following ImmPACT DAB Substrate Kit (Vector Laboratories, SK-4105) instructions. Composite images were taken on a Nikon NiE microscope under a 10× or 20× objective using a Nikon DS-Ri2 camera driven by Nikon Br Elements software. The percentage area of replacement fibrosis identified by fibronectin staining larger than a myocyte was quantified in entire cross-sections of the ventricles by an individual blinded to treatment using Adobe Photoshop CS6.

Echocardiography and cardiac strain analysis.

Endpoint echocardiography was performed on 3 and 9 month-old animals described above for use in flow cytometry analyses. Baseline and endpoint echocardiography was performed in Fiona/dko mice used for anti-CD3 or isotype control treatment described above. Longitudinal echocardiography was also performed in a separate cohort of Fiona/dko mice (n= 5M, 4F). Echocardiography was performed and analyzed by an investigator blinded to genotype and treatment in mice lightly anesthetized with 1.5% isoflurane. Measurements were taken using B-mode in the parasternal short axis view at the level of the papillary muscles with a Vevo3100 FUJIFILM VisualSonics system and MX550D transducer. To perform speckle tracking analysis, tracking points were placed along the endocardial and epicardial border of cine loops of parasternal short axis views using VevoStrain (Version 3.1.1), which allowed for frame-by-frame tracking through at least 3 cardiac cycles for calculations of strain and strain rates. M-mode images were created from the B-mode images and used to determine all other functional parameters by placing points along the endocardial border over at least 3 cardiac cycles for automatic calculation by the software.

RNA-sequencing.

Total RNA was isolated from frozen ventricular heart tissue from 9-month-old Fiona/dko (n = 2M, 1F) and Het/Fiona (n = 2M, 1F) mice after homogenization using TRIzol reagent (Life Technologies, Grand Island, NY, United States), according to the manufacturer’s instructions, followed by incubation with DNase I (RQ1; Promega, Madison, WI, United States). DNase-treated RNA concentration was determined spectrophotometrically and RNA integrity was measured using the Agilent 2100 Bioanalyzer (Agilent Technologies, Palo Alto, CA, United States). Additional quality control, library preparation and sequencing were performed by Novogene (Plant and Animal Eukaryotic mRNA-seq with reference, WBI-Quantification). Analysis to generate differential gene expression lists (using DEseq2, at padj < 0.1 and |log₂ fold change| ≥ 1) and gene ontology (GO) dot plots were done using Novogene NovoMagic software. To obtain a list of all “T cell” -related genes, GO-terms with “T cell” were extracted from all the GO terms downloaded from BioMart. The 172 distinct T cell-related terms obtained comprised 801 distinct genes. The overlapping genes between the total “T cell” related genes and the differentially expressed genes from the Fiona/dko vs Het/Fiona comparison were subsetted out. For the heatmaps, hierarchical clustering and plotting of normalized counts of the differentially expressed genes using the R-package pheatmap was performed. Normalization was performed across rows by calculating the z-score by subtracting the mean and then dividing by the standard deviation. Additional R-packages used for analysis include tidyverse and ggplot2.

Statistical Analysis.

Quantitative data are displayed as individual data points or for percent of CD45⁺ cells as mean ± SEM. All statistical analyses were performed using GraphPad Prism version 10.3.0 statistical software (GraphPad). The correct statistical test for each analysis was determined by first assessing the normality and variance of the data. For the flow cytometry data that was not normally distributed (cell numbers), values were log₁₀ transformed prior to conducting downstream statistical analysis and correlations. After transformation, a 1-way ANOVA followed by Dunnett’s T3 multiple comparison test was used to determine differences with age and genotype in flow cytometry data. For cardiac strain data comparison between genotypes, an unpaired Welch’s t test was performed. For strain correlations to flow data, a simple linear regression was performed using the log₁₀ transformed cell number values followed by analysis of covariance to test if the slopes of the regression lines were different between genotypes. Longitudinal echocardiography was analyzed by 2-way ANOVA with Tukey’s multiple comparisons test to determine changes over time and between genotypes. RNAseq data was analyzed by NovoMagic software with cutoff parameters of p_adj < 0.1 and |log₂ fold change| of ≥ 1. Treatment baseline to endpoint echocardiography was analyzed by linear regression analysis to track change within genotype over time and 2-way repeated measures ANOVA with uncorrected Fisher’s LSD multiple comparisons test was performed to determine differences between genotypes at each timepoint. For comparison between anti-CD3 antibody treated and isotype mice fibronectin staining, an unpaired Welch’s t test was performed.

Study approval.

Animal protocols were approved by the IACUC of The Ohio State University, in compliance with the NIH Guide for the Care and Use of Laboratory Animals (National Academies Press, 2011).

Data availability.

RNA sequencing data is available in NCBI-GEO accession number: GSE276884. Values for all data points in figures are reported in the Supporting Data Values file.

Results

Dystrophic cardiac inflammation is composed of T cells and myeloid cells

To specifically define the immune cell composition in hearts from the severe Fiona/dko model of DMD cardiomyopathy and the milder Het/Fiona littermates, we performed flow cytometry at the initial stage of cardiac damage at 3-months-of-age. We first performed flow cytometry using a similar myeloid panel as we had previously used to show that over 95% of the CD45⁺ immune cells in dystrophic quadriceps limb skeletal muscles are CD11b⁺ myeloid cells (28–30). The myeloid panel for dystrophic hearts included the pan-myeloid marker CD11b, the neutrophil marker LY6G, the macrophage/monocyte marker CD64, the reparative macrophage marker CD206, and the monocyte markers LY6C and CCR2 (Supplementary Figure S1). In contrast to dystrophic skeletal muscle, this panel showed that only 50.97 (± 7.94, n = 6) percent of CD45⁺ immune cells were represented by CD11b⁺ innate immunity myeloid cells in 3-month-old Fiona/dko hearts (Fig 1A, Table 1, Supplementary Table S1). Most of the CD11b⁺ myeloid cells were LY6G⁻ (78.68% ± 1.87), which included CD64⁺CD206⁺ macrophages (47.08% of CD11b⁺ ± 4.29) and CD64⁺LY6C^hi monocytes (35.10% of CD11b⁺ ± 2.08) (Table 1). Neutrophils made up 14.94 (± 2.13) percent of the CD11b⁺ myeloid cell population, or 7.44 (± 1.71) percent of the total CD45⁺ leukocytes (Table 1).

Figure 1. DMD cardiomyopathy is characterized by early lymphocyte infiltration.

(A) Representative flow cytometry plots of 3-month-old Fiona/dko mice with gating for CD45⁺:CD11b⁺ myeloid cells (top) and CD45⁺:CD3⁺ T cells (bottom). (B-C) Log₁₀ transformed cells per heart for each maker in the myeloid (B) and lymphoid (C) flow cytometry panels performed on hearts from 3- and 9-month-old Fiona/dko and Het/Fiona mice. Raw numbers were log₁₀ transformed to correct for normality and statistical significance was determined by 1-way ANOVA and Dunnett’s T3 multiple comparisons test. Myeloid populations were detected by CD11b (all myeloid cells); LY6G (neutrophils); CD64 (monocytes/macrophages); LY6C (monocytes); CD206 (reparative macrophages); and CCR2 (pro-inflammatory marker) (B) and lymphoid populations were detected by CD3 (all T cells); CD4⁺ (helper [Th] and regulatory T cells [Treg]); CD4⁻ (cytotoxic T-cells); FOXP3⁺ (regulatory T cells); FOXP3⁻ (conventional Th cells); and CD19 (B-cells) (C) 3-month Fiona/dko: n = 6; 3-month Het/Fiona: n = 7; 9-month Fiona/dko: n = 15; 9-month Het/Fiona: n = 15. * = p < 0.05; ** = p < 0.01, *** = p < 0.001.

Table 1.

Myeloid and lymphoid cell percentages of parent populations from 3-month-old Fiona/dko and Het/Fiona heart flow cytometry.

	Percent of CD45+ (%) Mean (± SEM)		Percent of CD11b+ (%) Mean (± SEM)		Percent of CD3+ (%) Mean (± SEM)
Myeloid Markers	Fiona/dko	Het/Fiona	Fiona/dko	Het/Fiona	Fiona/dko	Het/Fiona
CD11b+	50.97 (± 7.94)	38.27 (± 8.95)	-	-	-	-
CD11b+:LY6G−	40.44 (± 6.64)	29.74 (± 7.83)	78.68 (± 1.87)	74.69 (± 2.41)	-	-
LY6G−:CD64+CCR2+	2.90 (± 0.70)	2.88 (± 1.12)	5.51 (± 0.90)	6.00 (± 1.23)	-	-
LY6G−:CD64+CD206+	24.71 (± 5.08)	16.39 (± 5.82)	47.08 (± 4.29)	37.09 (± 5.04)	-	-
LY6G−:CD64+LY6Chi	18.50 (± 3.47)	15.64 (± 5.41)	35.10 (± 2.08)	35.35 (± 4.43)	-	-
LY6G−:CD64+LY6Clo	1.27 (± 0.42)	0.53 (± 0.18)	2.33 (± 0.57)	1.26 (± 0.25)	-	-
CD11b+:LY6G+	7.44 (± 1.71)	6.78 (± 0.95)	14.94 (± 2.13)	20.51 (± 2.51)	-	-
Lymphoid Markers	Fiona/dko	Het/Fiona	Fiona/dko	Het/Fiona	Fiona/dko	Het/Fiona
CD3+	30.69 (± 2.52)	34.77 (± 3.12)	-	-	-	-
CD3+:CD4+	20.79 (± 1.53)	22.14 (± 1.74)	-	-	68.44 (± 3.64)	64.66 (± 3.38)
CD4+:FOXP3+	6.16 (± 0.83)	7.16 (± 0.79)	-	-	20.40 (± 2.50)	20.61 (± 1.49)
CD4+:FOXP3−	14.63 (± 1.59)	14.94 (± 1.05)	-	-	48.03 (± 4.18)	43.94 (± 2.56)
CD3+:CD4−	9.90 (± 1.65)	12.62 (± 1.86)	-	-	31.56 (± 3.64)	35.34 (± 3.38)
CD19+	16.17 (± 4.20)	23.98 (± 5.53)	-	-	-	-

Percentage makeup of 3 parent populations (CD45, CD11b, and CD3) were calculated for each subpopulation as the mean ± SEM for 3-month-old Fiona/dko (n = 6) and Het/Fiona (n = 7) mice. Percentages were not significantly different between genotypes as determined by unpaired t-test with Welch's correction.

Since myeloid cells comprised only half of dystrophic heart immune cells, we next sought to determine the identity of the remainder of the immune cells. We performed flow cytometry with an additional panel that contained markers for lymphocytes, including the pan-T cell marker CD3, the helper T cell marker CD4, the regulatory T cell marker FOXP3, and the B-cell marker CD19. Surprisingly, this panel revealed that 3-month-old Fiona/dko hearts also contain 30.5 (± 2.55, n = 6) percent CD3⁺ T cells in contrast to dystrophic skeletal muscles (Fig 1A). Most of these CD3⁺ T cells were CD4⁺ (68.44% ± 3.64) (Table 1). In addition, CD19⁺ B-cells made up 16.17 (± 4.20) percent of the CD45⁺ leukocytes (Table 1). Percentages of myeloid or lymphoid cells in 3-month-old Fiona/dko were not significantly different from Het/Fiona littermates (Table 1).

To characterize how the immune profile changes throughout the disease process, we performed flow cytometry using these same myeloid and lymphoid panels on 9-month-old Fiona/dko and Het/Fiona mice and compared this data to the 3-month-old (mo) mice. We chose the 9-month timepoint because this is when ejection fraction is significantly decreased yet damage and fibrosis has plateaued in Fiona/dko mice (9), indicating the presence of a possible mechanism driving heart dysfunction. For all the myeloid cell markers measured, we surprisingly did not observe any significant differences between genotypes or ages (Fig 1B). Myeloid cells, including neutrophils (LY6G⁺) and macrophages/monocytes (CD64⁺), were still present chronically through 9-months-of-age in both models (Fig. 1B and Supplementary Table S1). However, there were significant differences in all the lymphocyte markers measured between ages but not genotypes (Fig 1C). From 3-months to 9-months within both Fiona/dko and Het/Fiona, lymphocytes significantly decreased (Fig 1C). Notably, CD3⁺ cells decreased from 38,904 (± 13,013) cells per heart to 4,364 (± 1,391) cells per heart from 3- to 9-months in Fiona/dko mice (Supplementary Table 1). To correct for normality, the cell per heart values were log transformed and statistical analysis was done on the transformed values (Fig 1B-C).

The persistence of both myeloid and lymphoid immune cells through 9-months-of-age demonstrates that dystrophic cardiomyopathy is characterized by chronic inflammation, while the significant difference in lymphoid immune cell numbers between the two ages indicates that much of the active inflammation occurs early in the disease course (Fig 1B-C). Since the presence of T cells is a recent finding in dystrophic hearts and dysregulation of CD4⁺ T cells has been implicated in other forms of cardiomyopathy and heart failure (35–37, 39, 41–44), we looked more specifically at two subsets of CD4⁺ T cells to identify whether they change between genotypes and age. These subsets included conventional Th cells (CD4⁺FOXP3⁻) and Treg cells (CD4⁺FOXP3⁺). Th and Treg cells were not significantly different between genotypes, but the most significant difference in lymphocyte number with age of Fiona/dko mice was in CD4⁺ helper T cells and both subpopulations of CD4⁺ cells: FOXP3⁺ and FOXP- (3mo vs. 9mo Fiona/dko: CD4⁺ p = 0.0002, CD4⁺FOXP3⁺ p = 0.0004, CD4⁺FOXP3⁻ p = 0.0002; 3mo Fiona/dko: n = 6, 9mo Fiona/dko: n = 15) (Fig 1C). This result could indicate that CD4⁺ helper T cells are playing an important role at the 3-month time point when much of the active, damaging inflammation is peaking. This data aligns with their usual role of responding to innate immune signals and secreting cytokines and chemokines to recruit other inflammatory cells (45, 46). Overall, this observation suggests that early treatment before inflammation peaks, and even substantially before function begins to decrease, may be important for preventing cardiomyopathy.

To verify whether 3-months-of-age is the earliest appearance of inflammation in our dystrophic cardiomyopathy mouse model, we performed immunohistochemistry to identify CD3⁺ cells on heart sections from 2-, 3-, 6-, 9- and 12-month-old Fiona/dko mice. We observed that a few isolated CD3⁺ T cells were present at 2-months-of-age but were not found in clusters nor were nearly as prevalent as observed in 3-month-old mice, where they first appeared as infiltrative clusters and then persisted throughout older ages, validating flow cytometry observations at 3-months-of-age (Fig 2A).

Figure 2. CD3⁺ T cell initial infiltration begins at 3-months-of-age and persists through 18-months but is prevented by μDystrophin gene therapy.

(A) Representative areas of one 2-month Fiona/dko heart, one Het/Fiona (labeled individually) and one C57 mouse (labeled individually) stained for CD3⁺ T cells (top row) showing no T-cell infiltration into the myocardium in wild-type mice or in Het/Fiona or Fiona/dko mice at 2 months-of-age. Representative images of CD3 staining of n=3 different hearts from Fiona/dko mice at ages 3, 6, 9 and 12 months of age show T-cell infiltration starts by 3 months and persists through 12 months (4 bottom rows). Scale bar: 100 μm. (B) Representative composites (top) and areas (bottom) of CD3 staining of 18-month-old Fiona/dko mice that were either untreated or treated at 4-weeks-of-age with (AAV)-micro-dystrophin (μDys5) gene therapy, respectively. μDys5 treated mice had qualitatively less CD3 infiltration than untreated mice. n = 5 mice/group; scale bars: 500 μm (top), 100 μm (bottom).

Since gene therapy delivery of micro-dystrophin has recently been FDA-approved, we stained heart sections from our previously published micro-dystrophin 5 (μDys5) gene therapy treated 18-month-old Fiona/dko mice to determine whether T cells are affected by this treatment (47). μDys5 treatment completely prevented pathology and functional decline in dystrophin Fiona/dko hearts and we previously showed that treatment prevented all immune cell (CD45⁺) infiltration (47). These 18 month-old μDys5 treated hearts also lacked clusters of CD3⁺ T cells and appeared similar to wild-type hearts, compared to clusters of infiltrated T cells present in untreated 18 month-old Fiona/dko mice (Fig 2B). Therefore, therapy that prevents dystrophic cardiac pathology and dysfunction also prevents T cell infiltration.

To start dissecting why Fiona/dko and Het/Fiona mice have similar numbers of immune cells but otherwise have significant phenotypic differences, including significantly different ejection fraction and fibronectin staining at 9-months-of-age (9), we investigated correlations of functional outcomes with immune cell numbers. We performed echocardiography on 3-month and 9-month-old Fiona/dko and Het/Fiona mice 1–2 days before they were dissected for flow cytometry. Measurements and calculations of whole heart functional parameters showed differences between Fiona/dko and Het/Fiona at both 3 and 9 months-of-age for stroke volume (p = 0.05, 3 months; p = 0.01, 9 months), ejection fraction (p = 0.03, 3 months; p = 0.007, 9 months) and fractional shortening (p = 0.04, 3 months; p = 0.007, 9 months), but no differences in other structural parameters at either age (Supplementary Table S2). Short axis B-mode images were analyzed by speckle tracking software to determine circumferential strain (CS) and circumferential strain rate (CSR) (Fig 3A) as previously described (47, 48). As these measurements are inherently negative, a reduced magnitude equates to reduced function. Overall, 9-month-old Fiona/dko mice had significantly decreased CS compared to Het/Fiona mice (p = 0.0006; Fiona/dko: n = 14, Het/Fiona: n = 12) (Fig 3B). When CS and CSR measurements were correlated with the number of CD3⁺, CD4⁺ and CD4⁻ cells per heart (log transformed), the linear regression analysis was divergent between Fiona/dko mice and Het/Fiona mice. Notably, Fiona/dko mice exhibited a negative correlation between strain (absolute number) and number of T cells (m = slope: CD3⁺: m = −3.352, CD4⁺: m = −2.550, CD4⁻: m = −3.592; Fiona/dko: n = 14, Het/Fiona: n = 12) (Fig 3C). The difference in slopes of the linear regression analysis between the genotypes was significantly different when correlating CS to the log of CD3⁺ cells per heart (p = 0.0288), CD4⁺ cells per heart (p = 0.0201) and CD4⁻ cells per heart (p = 0.0155) (Fig 3C). Thus, this data suggests a phenotypic difference between the T cells that may drive worsening function in severe dystrophic cardiomyopathy in the Fiona/dko model compared to less severe disease in Het/Fiona hearts, despite similar numbers of infiltrated T cells. To further investigate the timing of CS abnormalities observed at single time points in cohorts used for flow cytometry, we also performed longitudinal echocardiography CS measurements on a separate cohort of Fiona/dko mice at 3-, 6-, and 9-months-of-age, which showed similar significant worsening function (Fig 3D). The CS was significantly reduced in Fiona/dko mice from 3-months- to 9-months-of-age (p = 0.0470) (Fig 3D).

Figure 3. Fiona/dko mice have worse cardiac strain compared to Het littermates and exhibit a phenotypic divergence when correlated with T cells.

(A) Representative images of Fiona/dko (left) and Het Fiona (right) hearts from speckle tracking analysis with green lines indicating movement of the ventricle throughout the cardiac cycle and yellow dots indicating the location of the wall at diastole. (B) Circumferential strain (CS) of 9-month-old Fiona/dko and Het/Fiona mice demonstrated significantly worse function in the Fiona/dko mice. The Y-axis is inverted for easier visualization since a more negative number equals better strain. Statistical significance was determined by an unpaired t test with Welch’s correction. p = 0.0006; 9-month Fiona/dko: n = 14; 9-month Het/Fiona: n = 12. (C) Correlation of circumferential strain (CS, top) and circumferential strain rate (CSR, bottom) to log₁₀ transformed CD3⁺, CD4⁺ or CD4⁻ cells per heart for each 9-month-old Fiona/dko and Het/Fiona mouse from the flow cytometry analysis. The Y-axis and sign of the slopes (m ±) are inverted for easier visualization since a more negative number equals better strain. Statistical significance was determined by independently calculated simple linear regression analysis followed by analysis of covariance (ANCOVA) to test if the slopes of the two lines were significantly different. p-values are labeled on each individual graph and show that higher numbers of CD3⁺, CD4⁺ and CD4⁻ T cells correlate with worse circumferential strain only in Fiona/dko mice. Slope (m) is also shown only for Fiona/dko mice. 9-month Fiona/dko: n = 14; 9-month Het/Fiona: n = 12. (D) Longitudinal echocardiography at 3-, 6-, and 9-months-of-age on a separate cohort of Fiona/dko mice not used for flow cytometry. 9-month vs. 3-month: p = 0.0470; n = 5 M; 4F.

T cell related genes are significantly different in hearts of Fiona/dko versus Het/Fiona mice

Since T cell numbers weren’t different between genotypes yet were correlated with reduced function only in Fiona/dko, we next investigated whether whole heart gene expression varied between the two models. We hypothesized that the more severe dystrophic model, Fiona/dko, would have significantly increased pro-inflammatory and pro-fibrotic gene expression markers. Heart ventricles from 9-month-old Fiona/dko (n = 2M, 1F) and Het/Fiona (n = 2M, 1F) mice were dissected and RNA was extracted and sequenced. We used 9-month-old Fiona/dko and Het/Fiona mice since this is when left ventricular ejection fraction is significantly different from Het/Fiona and indicative of heart failure based on our previous studies (9, 47). There were 449 (p_adj < 0.1; |log₂foldchange| > 1) significantly differentially expressed genes (DEGs) in Fiona/dko versus Het/Fiona mice (Fig 4A, Supplementary Table S3). Notably, inflammatory and fibrosis-related genes were upregulated in Fiona/dko versus Het/Fiona (Fig 4B). We next searched for all gene ontology (GO) terms that contained “T cell” and extracted all the genes that fall within those GO terms, which gave us a list of 801 distinct genes. Then, we cross-referenced this T cell related gene list to our significant DEG list to identify T cell related genes that were differentially expressed between the two models.

Figure 4. Fiona/dko exhibit differential gene expression including T cell related genes.

(A) Volcano plot showing differential gene expression between 9-month-old Fiona/dko and Het/Fiona mice with the significance cutoff parameters of p_adj < 0.1 and |log₂ fold change| ≥ 1; n = 3 mice/group (B) Gene ontology dot plot for all significantly upregulated genes expressed in 9-month-old Fiona/dko compared to Het/Fiona mice. n = 3 mice/group. (C) Heat map depicting the 68 significantly differentially expressed T cell related genes (p_adj < 0.1) between 9-month-old Het/Fiona and Fiona/dko mice. These genes were cross-referenced with a list of all genes pulled from a search for any gene ontology terms containing “T cell”. Normalized counts are plotted.

From this comparison, we identified 68 T cell related genes that were significantly different (p_adj < 0.1) between Fiona/dko and Het/Fiona, with 51 being increased in Fiona/dko (Fig 4C; Supplementary Table S4). Some of the upregulated genes with the largest fold changes included galectin-3 which is involved in cell adhesion and T cell regulation, MALT1 that plays a role in CD4 Th17 activation and cytokine production, IL7 and its receptor IL7R, which are involved in T cell homeostasis and survival, and CD44 which is a marker of activated T cells (Supplementary Table S4) (49–53). Conversely, there are some anti-inflammatory gene changes such as upregulated annexin A1 that plays a role in glucocorticoid anti-inflammatory mechanism and NLRC3 which inhibits NF-κB and T cell activation (54–56). It is possible that Tregs may be trying to limit inflammation but are ultimately ineffective. However, annexin A1 has also been shown to skew T cell differentiation towards pro-inflammatory Th1 as opposed to anti-inflammatory Th2 (57). Thus, the Fiona/dko heart microenvironment is exhibiting a differential inflammatory phenotype and functional reactivity related to T cell signaling that may explain their worse pathology and progressive heart dysfunction despite stable fibrosis.

Systemic T- cell depletion reduces onset of pathology in Fiona/dko hearts

Since cardiac damage in utrophin/dystrophin-deficient hearts begins at approximately 8-weeks-of-age (58) and cardiac inflammation appears to peak at 3-months-of-age with more severe pathology, we next investigated the contribution of T cells to this early pathological stage. We performed a power calculation (alpha 0.05, power 0.8) based on our previously published data on the Fiona/dko model that predicts a sample size of groups of n=3 with an actual power estimated to equal 0.97 to distinguish differences in fibronectin accumulation indicative of fibrosis, since the fibrosis is already so dramatic by 3 months. We systemically depleted T cells by injecting 8-week-old Fiona/dko mice with mouse anti-CD3 neutralizing antibody (n = 2M, 1F) or an isotype control antibody (n = 3M) for 5 weeks until 3-months-of-age and then dissected the mice for histology (Fig 5A). Before and after the treatment, we performed echocardiography and flow cytometry of blood. As expected, there was an observable decrease in circulating blood CD3⁺ T cells, including both CD4⁺ and CD8⁺ cells, in the anti-CD3 treated mice compared to isotype after the 5 weeks (Fig 5B). Circulating CD3⁺ T-cells composed 24.6 ± 3.3% of circulating CD45⁺ immune cells in the isotype control treated Fiona/dko mice compared to 2.8 ± 0.4% in the anti-CD3 treated Fiona/dko mice, further validating systemic depletion of CD3⁺ T cells with anti-CD antibody treatment.

Figure 5. CD3 neutralizing antibody treatment depletes circulating CD4⁺ and CD8⁺ T cells and is not detrimental to heart function.

(A) 5-week treatment timeline schematic. 8-week-old Fiona/dko mice were initially injected intraperitoneally (i.p.) with 200 μg of an anti-CD3 antibody or isotype control antibody once a day for the first 3 days. For the next 4 weeks, the mice were injected i.p. twice a week with 150 μg antibody or isotype. (B) Representative flow cytometry dot plots of blood leukocytes from anti-CD3 treated (top) or isotype (bottom) mice. Gating for CD45⁺:CD3⁺ is shown on the left, within this gate CD4⁺ and CD8⁺ gates were set as shown on the right. CD3⁺, CD4⁺ and CD8⁺ were substantially depleted in the treatment group. n = 3 mice/group. (C) Echocardiographic strain analysis of isotype and anti-CD3 treatment mice. Circumferential strain (CS) and circumferential strain rate (CSR) were not significantly different between the groups before or after treatment (top), but anti-CD3 treated mice trended positively over the course of the treatment while isotype mice trended negatively by linear regression analysis (bottom; m = slope). The Y-axis and sign of the slopes (± m) are inverted for easier visualization since a more negative number equals better strain. Statistical significance was determined by 2-way repeated measures ANOVA with uncorrected Fisher’s LSD multiple comparisons test.

Since this study was designed to investigate onset of pathology and not powered to detect functional differences which are minimal and not indicative of heart failure at this stage, CS and CSR were not yet significantly different with treatment although the anti-CD3 treated mice trended positively (CS: m = 0.695) while isotype trended negatively (CS: m = −0.103) over time (Fig. 5C). This data supports that T cell depletion does not have any negative impact on cardiac function. Other echocardiography measurements calculated from short axis m-mode were also not yet significantly different (Supplementary Table S5).

The primary outcomes of this study, histopathology and fibrosis were improved by T cell depletion (Fig 6A-E). H&E and CD3 histology showed a decrease in pathology and T cell infiltration, respectively, in treated Fiona/dko mice compared to isotype-treated controls. These data confirmed that systemic CD3 depletion resulted in reduced T cell localization in treated hearts (Fig. 6A-B). In both treated and untreated hearts, all immune cell staining (CD45 leukocyte, CD3 T cell and CD11b myeloid) colocalized to any remaining areas of damage, although treated hearts had qualitatively less observable staining by all markers overall (Fig 6A-B). The overlap of CD11b and CD3 staining indicates that myeloid and T cells are likely communicating to infiltrate tissue and alter the pathology.

Figure 6. CD3 depletion improves pathology and prevents myeloid cell infiltration in Fiona/dko hearts.

(A) Microscope composite images of hearts from anti-CD3 neutralizing antibody treated (anti-CD3 tx) and isotype injected Fiona/dko mice stained with (from top to bottom): hematoxylin and eosin (H&E), CD45 total leukocytes, CD3 T cells, CD11b myeloid (Mye) cells. n = 3 mice/group, scale bars: 500 μm. (B) The same area of heart sections stained independently for CD45 total leukocytes, CD3 T cells and CD11b myeloid cells (Mye) showing colocalization of immune cells within areas of cardiac damage. The anti-CD3 treated mouse (top) shows qualitatively less infiltration even in an area containing some immune cells that is much rarer than the isotype mouse (bottom); Scale bar: 100 μm. (C) Composite images of anti-CD3 treated and isotype treated hearts from Fiona/dko mice stained for fibronectin, a marker of fibrotic scarring; Scale bar: 500 μm. (D) Co-immunostaining for fibronectin and IgG infiltration within the myocardium in anti-CD3 treated and isotype control Fiona/dko hearts staining in dystrophic hearts. Outside of vessels in damaged hearts, IgG leaks into areas of fibrosis and also is taken up into damaged cardiomocytes. IgG is observed overlapping with fibronectin only in the isotype controls and is not present within cardiomyocytes in either group showing that there are no quantifiable numbers of dying cardiomyocytes at this time-point since most damage occurs prior to 3 months-of-age; Scale bar: 100 μm. (E) Quantification of the fibronectin staining of entire cross-sections of ventricular tissue shown in (C) completed by a researcher blinded to treatment. Values are shown as percent of total area of the ventricular tissue, with the isotype Fiona/dko mice having significantly higher percentage of fibrosis. Statistical significance was determined by an unpaired t test with Welch’s correction. n = 3 mice/group; p = 0.0277.

Anti-CD3 treated Fiona/dko reduced the observed area of fibrotic scarring within the ventricles as determined by fibronectin immunostaining (Fig 6C), which we have previous demonstrated to be the most sensitive marker of dystrophic cardiac fibrosis (9, 47, 59). We also stained for the presence of endogenous IgG staining within cardiomyocytes, which we have previously demonstrated to be a sensitive assay for identifying dystrophic cardiomyocytes with damaged membranes as they start degenerating (60–63). As cardiac enzymes leak into the serum, serum proteins including IgG accumulate into the compromised cardiomyocytes and result in bright staining. IgG also shows diffuse staining in inflamed and fibrotic areas. Only diffuse IgG staining that overlapped with fibronectin staining was present only in isotype controls, but no IgG localization was present outside of blood vessels in anti-CD3 treated hearts (Fig 6D). This data suggests that no dying cardiomyocytes are present at this time-point in any of the mice since most myocardial damage occurs prior to 3 months-of-age (58). Quantification of the area of fibronectin localization demonstrated significantly less fibrosis in the anti-CD3 treated Fiona/dko hearts compared to isotype controls (p = 0.0277; n = 3 mice/group) (Fig 6E). Thus, whole body depletion of T cells in the Fiona/dko severe model of dystrophic cardiomyopathy was effective at improving histopathology onset in the heart.

Discussion

Duchenne muscular dystrophy (DMD) cardiomyopathy is characterized by chronic inflammation that is different from that seen in the skeletal muscle, thus requiring development of unique therapies to prevent progression to heart failure, which is the leading cause of death for DMD patients. Currently, corticosteroids are used to manage inflammation with some benefit but come with numerous side effects (26). Multiple immune cells are known to play a role in DMD skeletal muscle pathology (20, 64, 65), but less is known about the inflammatory microenvironment of the heart. Previously we demonstrated that mdx mouse skeletal muscle inflammatory milieu contained over 90% myeloid cells (28–30). In this study, we demonstrate T cells as a key contributor to DMD cardiomyopathy pathogenesis. Our results corroborate and expand on the recent findings of Morroni et al. that an inhibitor of PKCƟ, a protein involved in T cell activation, was able to reduce all investigated immune cell infiltration, necrosis and fibrosis in the heart in exercised mdx mice, although this model does not show contractile dysfunction indicative of heart failure (34).

In the Fiona/dko model of isolated DMD cardiomyopathy, T cells comprise a large portion of the immune cell population in contrast to what we’ve previously observed in skeletal muscles of mdx and het mice. Furthermore, T cell numbers assessed by both flow cytometry and histology peaked at 3-months-of-age, which is early in the disease progression when much of the active damage and fibrosis is occurring, indicating that T cells may be playing a role in driving these processes. T cells then persisted through 18-months-of-age as seen on histology, highlighting the chronic inflammatory nature of the disease. This data, along with the differential expression between Fiona/dko and Het/Fiona of genes related to T-cell signaling, indicates that T cells may also be important in driving the heart dysfunction seen at this later stage of disease, which cannot be explained by fibrosis that plateaus and remains stable after 6-months-of-age in these mice (9). Indeed, T cells and more specifically dysregulated CD4⁺ T cells subtypes have been implicated in several other types of cardiomyopathy (36, 37, 39, 41–44) and may even be playing a role in the myocarditis observed in DMD gene therapy patients (35, 66, 67).

We examined 2 subtypes of CD4⁺ helper T cells in this study: conventional Th cells (CD4⁺FOXP3⁻) and Treg cells (CD4⁺FOXP3⁺). Persistent Tregs have been shown to become dysregulated and contribute to ischemic cardiomyopathy pathology (68). In Fiona/dko mice, these regulatory cells follow the same trend as the other T cells, with an early peak of infiltration followed by significant decline in numbers but a chronic presence. Whether these cells become dysregulated during persistent inflammation as in ischemic cardiomyopathy will need to be investigated. A limitation of this study is that we did not perform flow cytometry on wild-type mouse hearts. However, a comprehensive study using multiple models and techniques has previously been conducted on wild-type mouse hearts demonstrating that 81.4 ± 1.4% of CD45⁺ immune cells are CD11b⁺ myeloid cells, and 3.1 ± 0.4% of CD45⁺ cells are CD3⁺ T cells. (69). Thus, the composition of infiltrating immune cells in dystrophic hearts includes a much higher number and proportion of T cells than observed in wild-type mice, which we confirm directly by immunostaining (Fig. 2). Future work should also further delineate the specific subpopulations of these CD4⁺ cells to determine their exact role and whether they are driving a pro-inflammatory environment. Overall, immunopathology peaks very early in the disease process, before cardiac dysfunction is detectable by strain or ejection fraction. For patients, this data suggests that therapeutic intervention may be necessary much earlier than currently prescribed, probably before heart dysfunction even at the level of myocardial strain abnormalities may be clinically detectable.

The similar numbers of heart immune cells via flow cytometry between Fiona/dko and Het/Fiona were somewhat surprising. However, Fiona/dko hearts and the T cells infiltrating them had distinct phenotypic differences from that of the Het/Fiona mice. Fiona/dko had significantly decreased cardiac strain that correlated with increasing numbers of T cells and distinct changes in T cell gene expression pathways that could provide an explanation for how T cells may be driving the worsening function seen in Fiona/dko hearts compared to Het/Fionas. In addition to gene changes noted in the results, other notable upregulated genes with low p_adj and high fold change included midkine that is involved in repression of Tregs and is produced by other inflammatory CD4⁺ T cells, CXCL16 which is a chemoattractant of activated T cells, and beta-2 microglobulin which is a component of MHC I that presents antigen to CD8⁺ T cells (70–72). Thus, it appears that although the number of T cells weren’t significantly different between the two models, the more severe Fiona/dko does appear to exhibit more pro-inflammatory signaling, specifically including T cell signaling pathways. This data could explain the more severe pathology and functional decline seen in this model. More work will need to be done to differentiate whether this signaling is coming from the T cells or the cardiomyocytes or myeloid cells and whether this T cell signaling is driving damage or simply a result of ongoing damage. Future work will also need to explore how T cells interact with or modify the activity other immune cells, since myeloid cells are also still chronically present in the model and were shown to colocalize with CD3⁺ T cells.

Here, we present T cells as an important contributor to pathogenesis in a mouse model of isolated DMD cardiomyopathy. Although inhibiting all T cells in humans will not be a viable therapeutic strategy, this opens the door for more specific development of therapies targeting T cell signaling. This type of therapeutic approach may also be beneficial for patients with immune reactions to adeno-associated virus (AAV)-micro-dystrophin gene therapies. Future studies on T cell depletion and T cell signaling disruption will need to be conducted beyond 9-months of age in this model to assess effects on function.

Immunomodulatory treatments have shown promising results but also highlight the need for continued understanding of the inflammatory pathology of the heart and more targeted therapies. Furthermore, while gene therapies using recombinant AAV vectors have yielded positive results, some patients are experiencing a myocarditis-like reaction to the therapy that also warrants concurrent treatment with anti-inflammatories (67). Therefore, supplementing gene-based therapies with immunosuppression or immunomodulation will be important, as inflammation is a predominant feature when DMD is diagnosed, and gene therapy alone will not adequately prevent all immunopathology.

Supplementary Material

Supplementary Figure S1: 10.6084/m9.figshare.30099709

Supplementary Table S1: 10.6084/m9.figshare.30099805

Supplementary Table S2: 10.6084/m9.figshare.30099883

Supplementary Table S3. 10.6084/m9.figshare.30099889

Supplementary Table S4: 10.6084/m9.figshare.30099892

Supplementary Table S5: 10.6084/m9.figshare.30099904