Calcium handling dysfunction and cardiac damage following acute ventricular preload challenge in the dystrophin-deficient mouse heart

Abstract

Duchenne muscular dystrophy (DMD) is the most common muscular dystrophy and is caused by mutations in the dystrophin gene. Dystrophin deficiency is associated with structural and functional changes of the muscle cell sarcolemma and/or stretch-induced ion channel activation. In this investigation, we use mice with transgenic cardiomyocyte-specific expression of the GCaMP6f Ca²⁺ indicator to test the hypothesis that dystrophin deficiency leads to cardiomyocyte Ca²⁺ handling abnormalities following preload challenge. α-MHC-MerCreMer-GCaMP6f transgenic mice were developed on both a wild-type (WT) or dystrophic (Dmd^mdx-4Cv) background. Isolated hearts of 3–7-mo male mice were perfused in unloaded Langendorff mode (0 mmHg) and working heart mode (preload = 20 mmHg). Following a 30-min preload challenge, hearts were perfused in unloaded Langendorff mode with 40 μM blebbistatin, and GCaMP6f was imaged using confocal fluorescence microscopy. Incidence of premature ventricular complexes (PVCs) was monitored before and following preload elevation at 20 mmHg. Hearts of both wild-type and dystrophic mice exhibited similar left ventricular contractile function. Following preload challenge, dystrophic hearts exhibited a reduction in GCaMP6f-positive cardiomyocytes and an increase in number of cardiomyocytes exhibiting Ca²⁺ waves/overload. Incidence of cardiac arrhythmias was low in both wild-type and dystrophic hearts during unloaded Langendorff mode. However, after preload elevation to 20-mmHg hearts of dystrophic mice exhibited an increased incidence of PVCs compared with hearts of wild-type mice. In conclusion, these data indicate susceptibility to preload-induced Ca²⁺ overload, ventricular damage, and ventricular dysfunction in male Dmd^mdx-4Cv hearts. Our data support the hypothesis that cardiomyocyte Ca²⁺ overload underlies cardiac dysfunction in muscular dystrophy.

NEW & NOTEWORTHY The mechanisms of cardiac disease progression in muscular dystrophy are complex and poorly understood. Using a transgenic mouse model with cardiomyocyte-specific expression of the GCaMP6f Ca²⁺ indicator, the present study provides further support for the Ca²⁺-overload hypothesis of disease progression and ventricular arrhythmogenesis in muscular dystrophy.

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INTRODUCTION

Duchenne muscular dystrophy (DMD) is an X-linked recessive disorder caused by mutations in the gene for dystrophin and affects ∼1 in 5,000 live male births (1). Approximately 60% of dystrophin mutations are large insertions or deletions that lead to frameshift errors, whereas ∼40% are point mutations or small frameshift rearrangements (2). In most cases, dystrophin mutations lead to loss of functional dystrophin protein and an impairment in sarcolemma integrity (3). Cardiomyopathy is a leading cause of death in patients with DMD, necessitating additional information on the pathophysiology of DMD in cardiac tissue (4).

In striated muscle, dystrophin is part of the dystrophin-associated protein complex (DAPC). This complex involves the sarcolemma, cytoskeleton, signaling proteins, and scaffolding proteins. Ion channels and transporters are also established partners of the DAPC and include the plasma membrane Ca²⁺ ATPase (PMCA), Na⁺ channels, K⁺ channels, L-type Ca²⁺ channels, the Na⁺-Ca²⁺ exchanger (NCX), and numerous stretch-activated channels (SACs, including transient receptor potential (TRP) channels; 1, 5–10). In patients with DMD, as well as in animal models of dystrophin deficiency, the lack of dystrophin results in excessive stress and/or mechanical stretch-induced activity of Ca²⁺ influx pathways in both skeletal and cardiac muscle (6, 10–16). A prevailing hypothesis to explain DMD disease progression is one of dystrophin deficiency contributing to abnormal sarcolemmal Ca²⁺ flux balance, including abnormal sarcolemmal Ca²⁺ entry through Ca²⁺ channels, reverse mode NCX activity, and/or microtears of the sarcolemma (17). Ca²⁺-cycling defects can also arise from increased Ca²⁺ “leak” through the ryanodine receptor (RyR), reduced activity of the sarcoplasmic reticulum Ca²⁺ ATPase, excessive reactive oxygen species (ROS) production, mitochondrial dysfunction, and a complex vicious cycle that leads to cytosolic Ca²⁺ overload, mitochondrial permeability transition, activation of proteases, myocyte cell death, and progressive muscle damage (18–22). With advanced DMD progression, Ca²⁺-handling proteins change expression and/or function, such as decreased levels of RyR, decreased SR luminal Ca²⁺-binding proteins, decreased cardiac sarcoplasmic reticulum Ca²⁺ ATPase-2 expression, and an increase in resting [Ca²⁺]_i (23–26). However, given the complicated nature of disease progression in DMD, it remains unclear how susceptible the dystrophic myocardium is to acute preload (i.e., stretch) challenge. In this study, we use cardiomyocyte-specific expression of the GCaMP6f Ca²⁺ indicator in wild-type and dystrophic (Dmd^mdx-4Cv) hearts to test the hypothesis that acute ventricular preload challenge leads to Ca²⁺ handling abnormalities and cardiac dysfunction in the dystrophic heart.

METHODS

Genetic Animal Models

Animal procedures were approved by the Animal Care and Use Committee at the University of Missouri (Approval Reference No. 9581) and complied with all United States regulations involving animal experiments. Wild-type background (WT) or dystrophic Dmd^mdx-4Cv (Jackson laboratory Strain No. 002378) male mice at 3–7 mo of age were used to investigate the role of dystrophin deficiency on functional parameters. One 8-mo WT animal was included as an additional control experiment. Male mice were used in this investigation because of the X-linked inheritance of muscular dystrophy and potential translational relevance to human patients. Calcium measurements were performed in mice with cardiomyocyte-specific expression of the GCaMP6f Ca²⁺ sensor. For this purpose, Ai95D transgenic mice [Ai95(RCL-GCaMP6f)-D, Jackson Laboratory Strain No. 028865] were crossed with a-MHC-MerCreMer transgenic mice (Jackson Laboratory Strain No. 005657) and Dmd^mdx-4Cv mice to obtain α-MHC-MerCreMer × Ai95D GCaMP6f mice without (wild-type background, WT) and with the Dmd^mdx-4Cv allele (dystrophic). Tamoxifen injection (intraperitoneal injection of 1 mg/day for 5 days) induced uniform expression of the GCaMP6f Ca²⁺ sensor within cardiomyocytes as in previous studies (27). Mice were studied a minimum of 30 days post-tamoxifen injection to avoid adverse effects associated with activation of the Cre recombinase (28). Additional control experiments (cardiac pressure development studies) were performed in tamoxifen-treated wild-type and dystrophic mice with single transgenes associated with the breeding strategy (α-MHC-MerCreMer or Ai95D GCaMP6f).

Preload Challenge of Isolated Working Hearts

Modified Krebs–Henseleit buffer (KHB) was used as physiological saline for working heart experiments and contained the following: (in mM) 117 NaCl, 4.7 KCl, 1.2 MgSO₄, 1.2 KH₂PO₄, 25 NaHCO₃, 11.11 glucose, 0.4 caprylic acid, 1 pyruvate, 0.0023 Na-EDTA, and 1.8 CaCl₂. Mice were anesthetized with an intraperitoneal injection of ketamine-xylazine (100 mg/kg-5 mg/kg), and hearts were rapidly (∼30 s) excised and cannulated (within 3 min) via the aorta for retrograde coronary perfusion of oxygenated (95% O₂-5% CO₂) KHB at a constant afterload of 60 mmHg at 37°C. The pulmonary vasculature was ligated, and incisions were made in the right atrium and pulmonary artery to drain fluid and stabilize pressure. An incision was also made in the left atrium, and the atrium was cannulated with a custom-fabricated atrial cannula connected to a gravity-fed, temperature-controlled glass oxygenator (Radnoti, Covina, CA) to provide constant-pressure control of left ventricular preload (working heart mode). A 1.0-Fr tip pressure catheter was inserted into the left ventricle via the apex of the heart to monitor pressure development. After aortic cannulation, coronary effluent was collected for subsequent detection of lactate dehydrogenase (LDH) using the Promega luminescent LDH-Glo cytotoxicity assay kit following manufacturer protocol (Promega, Madison, WI). For pressure measurements, the right atrium was electrically stimulated at 6 Hz using a pair of platinum electrodes (voltage set, ∼50% higher than capture threshold, typically 4–5 V), and each heart was allowed to equilibrate in unloaded Langendorff mode for >5 min to obtain unloaded baseline functional measurements. The left ventricle was then subjected to elevated preload of 20 mmHg for 30 min via the atrial cannula. We chose this pressure for ventricular preload as this pressure results in the near-maximal sarcomere lengthening in mouse hearts (29). Following preload challenge, the heart was returned to unloaded Langendorff mode for a repeat collection of coronary effluent to assess poststretch LDH levels. Steady-state maximum pressure, rate of pressure development, rate of pressure decay, and heart rate were analyzed over 5–15-s intervals during three phases: 1) baseline Langendorff, 2) 20-mmHg initial maximum (defined as maximum pressure response following 20-mmHg preload elevation), and 3) 30 min post-20-mmHg preload elevation. Maximum rate of pressure development (dP/dt_max) and minimum rate of pressure decay (dP/dt_min) associated with each pressure response were analyzed using Labchart/Powerlab 8.1 software (AD Instruments). A baseline (Langendorff mode) left ventricular peak pressure of 40 mmHg or greater confirmed a successful physiological preparation and was used as an a priori inclusion criterion.

High-Speed, High-Resolution Confocal Imaging of GCaMP6f

Following preload challenge protocol, GCaMP6f-positive hearts were removed from the working heart apparatus and perfused via the aorta in unloaded Langendorff mode with the myosin II inhibitor blebbistatin (40 µm) to paralyze the heart for confocal imaging. A 1.5-mm-length segment of polyethylene tubing was inserted in the left ventricle via the mitral valve to facilitate drainage of venous coronary effluent from the chamber during blebbistatin-induced cardioplegia. The conversion from working heart mode to Langendorff mode interrupted aortic perfusion for <3 s, and once in Langendorff mode, the heart was continually perfused with oxygenated KHB. The heart was mounted on a custom-imaging platform and transported to an adjacent room to an inverted Leica SP5 confocal microscope for laser-scanning confocal fluorescence imaging of the subepicardium (×20 objective; 488-nm excitation, 500–540-nm emission; 27). Confocal imaging was performed at 25°C, with inclusion criteria of a sinus heart rate of >2 Hz or with constant rate stimulation of 3 Hz (electrical field stimulation). GCaMP6f-positive pixels and percent Ca²⁺-overloaded cells were assessed with a 512 × 512 pixel frame at 0.38 mm/pixel and 30 frames/s. Ca²⁺ transients were assessed with a 512 × 130 pixel frame at 0.38 mm/pixel and 83 frames/s. Percent GCaMP6f positive pixels were analyzed by an observer blinded to genotype and intervention using ImageJ software. A threshold was set for each image, and the image was converted to a binary for determination of the number of positive pixels per field of view (FOV).

Electrocardiogram Measurements

For assessment of cardiac arrhythmia, separate cohorts of (non-GCaMP6f) wild-type and Dmd^mdx-4Cv dystrophic mice were perfused in unloaded Langendorff mode to obtain baseline arrhythmia incidence, followed by elevated preload challenge for 30 min as described earlier. For this set of experiments, hearts were allowed to beat at sinus rhythm to avoid electrical stimulation-induced entrainment of pacemakers and/or overdrive suppression of arrhythmias. Average heart rate was determined over 10 s of steady-state pressure waveforms. Cardiac electrocardiograms (ECGs) were monitored continuously at baseline and throughout the preload challenge protocol using MLA1213 needle electrodes, with a set of three 1.5-mm shrouded socket monopolar electrodes (29 gauge) in conjunction with a FE231 Bio Amp and LabChart/Powerlab 8.1 software (AD Instruments). The position of the electrodes was adjusted for each experiment to obtain a distinct P wave and QRS complex for arrhythmia analysis. Cardiac arrhythmia incidence was assessed in the last 5 min of baseline and last 5 min of preload challenge at 20 mmHg. Premature ventricular complexes (PVCs) were defined as a premature ventricular ECG waveform preceding and/or independent of the atrial P wave. QRS complexes associated with PVCs were typically of higher amplitude, longer in duration, and of a distinct waveform versus sinus driven QRS complexes. In each minute of analysis, the presence of arrhythmias was quantified using a 0, 1, and 2 arrhythmia score system, with score of 0 representing no arrhythmias, a score of 1 corresponding to isolated PVCs, and a score of 2 corresponding to salvos of PVCs > 3 or ventricular tachycardia/fibrillation. The 5-min average arrhythmia score per heart was used for final data analysis.

Data Analysis and Statistics

All data are presented as individual observations and sample means (x) ± SE. Minimum sample sizes were determined by an a priori power analysis (power = 0.80). Summary data were analyzed using one-tailed Student’s t test [homoscedastic (F test of variance not different) or heteroscedastic (F test of variance significantly different) as appropriate], or Mann–Whitney test for nonparametric samples or as appropriate to experimental design and indicated by figure legend. Data were analyzed using Spark AN 5.5.4.0, LabChart 8.1.16, and GraphPad 10.0.2. P < 0.05, P < 0.01, P < 0.001, and P < 0.0001 indicate statistically significant differences.

Blinding

For GCaMP6f experiments, the experimentalist was not blinded to genotype to ensure correct animal model (i.e., GCaMP6f positive vs. GCaMP6f negative animal) entered the confocal imaging protocol. For non-GCaMP6f electrocardiogram studies the experimentalist was blinded to genotype during both data acquisition and analysis, with genotype of animal (WT or Dmd^mdx-4Cv) determined by the University of Missouri Animal Modeling Core after completion of experimental studies and data analysis.

RESULTS

Left ventricular pressure was monitored in isolated, perfused hearts of 3- to 7-mo-old wild-type (WT) or dystrophic Dmd^mdx-4Cv GCaMP6f mice (Fig. 1) subjected to sustained left ventricular preload challenge. In the absence of ventricular preload (unloaded Langendorff mode), hearts of both wild-type and dystrophic mice exhibited similar maximum systolic pressure (Fig. 2, A–C), rate of pressure development (Fig. 2D), and rate of pressure decay (Fig. 2E). Following preload elevation to 20 mmHg, both wild-type and dystrophic hearts exhibited a similar augmentation in systolic maximum pressure (Fig. 2F), rate of pressure development (Fig. 2G), and rate of pressure decay (Fig. 2H). Maximum pressure (Fig. 3A), rate of pressure development (Fig. 3B), and rate of pressure decay (Fig. 3C) were also similar between wild-type and dystrophic hearts at the conclusion of the 30-min sustained preload challenge. Lactate dehydrogenase (LDH) release from coronary effluent was similar between wild-type and dystrophic hearts under unloaded Langendorff conditions (Fig. 4A). However, LDH was significantly elevated in dystrophic versus wild-type hearts following the preload challenge protocol (Fig. 4B).

Figure 1.

Triple-transgenic MerCreMer × Ai95 (GCaMP6f) × Dmd^mdx-4Cv transgenic construct. Transgenic unaffected (wild type, WT; A) or Dmd^mdx-4Cv dystrophic mouse (X chromosome; B) contains the α-MHC-MerCreMer (MCM) transgene located on chromosome 19 and the Rosa-loxP-STOP-loxP-GCaMP6f transgene located on chromosome 6. C: when mice are treated with tamoxifen, Cre-mediated excision of the floxed STOP cassette leads to expression of the GCaMP6f Ca²⁺ sensor within cardiomyocytes. Images created using a licensed version of BioRender.com.

Figure 2.

Cardiac function in ex vivo perfused hearts. Example trances of left ventricular pressure (top) and rate of pressure change (dP/dt, bottom) in hearts from wild-type (WT; A) and Dmd^mdx-4Cv (B) mice at 0-mmHg (left) and 20-mmHg preload (right). Summary data of maximum pressure development (P_max, C), maximum rate of pressure development (dP/dt_max, D), and relaxation rate (dP/dt_min, E) of WT (circles) and Dmd^mdx-4Cv (squares) hearts at 0-mmHg preload. Summary data of P_max (F), dP/dt_max (G), and dP/dt_min (H) of WT (circles) and Dmd^mdx-4Cv (squares) hearts at 20-mmHg preload. Data include hearts with (green symbols) and without (open symbols) cardiomyocyte GCaMP6f expression. P > 0.05 for all comparisons; not significant (ns); WT vs. Dmd^mdx-4Cv, unpaired t test. n = 6, 3–6-mo WT and n = 7, 4–7-mo Dmd^mdx-4Cv hearts.

Figure 3.

Cardiac function following sustained preload challenge. Summary data of maximum pressure development (P_max, A), maximum rate of pressure development (dP/dt_max, B), and relaxation rate (dP/dt_min, C) of wild-type (WT, circles) and Dmd^mdx-4Cv (squares) hearts following 30 min of 20-mmHg preload challenge. Data include hearts with (green symbols) and without (open symbols) cardiomyocyte GCaMP6f expression. P > 0.05 for all comparisons; not significant (ns); WT vs. Dmd^mdx-4Cv, unpaired t test. n = 6, 3–6-mo WT and n = 7, 4–7-mo Dmd^mdx-4Cv hearts.

Figure 4.

Cardiac damage in dystrophic hearts following sustained preload challenge. Summary data of coronary lactate dehydrogenase (LDH) in wild-type (WT, circles) and Dmd^mdx-4Cv (squares) hearts at 0-mmHg (A) and 30 min following 20-mmHg preload challenge (B). Data include hearts with (green symbols) and without (open symbols) cardiomyocyte GCaMP6f expression. ***P = 0.0007; WT vs. Dmd^mdx-4Cv, one-tailed Mann–Whitney U test. n = 6, 3–6-mo WT and n = 8, 3–6-mo Dmd^mdx-4Cv hearts.

To visualize cardiomyocyte GCaMP6f signal and Ca²⁺ handling abnormalities following the 30-min preload challenge, hearts were perfused under unloaded Langendorff conditions with the myosin II inhibitor blebbistatin and imaged using laser scanning confocal fluorescence microscopy. Hearts of dystrophic, but not wild-type, GCaMP6f mice exhibited regions devoid of fluorescence signal (Fig. 5, A and B). Quantification of GCaMP6f positive pixels within the field of view indicated a significant decrease in the percentage (%) of GCaMP6f positive pixels in dystrophic hearts consistent with myocyte dysfunction and/or damage (Fig. 5C). Control experiments in dystrophic hearts in the absence of preload confirmed GCaMP6f fluorescence was uniform across the myocardium in this mouse model and not because of disease progression alone at this age in mice (Fig. 5A). Furthermore, in dystrophic hearts following the preload challenge, cardiomyocytes with residual GCaMP6f fluorescence frequently exhibited Ca²⁺ overload and/or nonsteady-state Ca²⁺ signals including Ca²⁺ waves (Fig. 6, A–C).

Figure 5.

Loss of GCaMP6f-positive cardiomyocytes in dystrophic hearts following sustained preload challenge. A: confocal images of subepicardial cardiomyocytes in wild-type (WT) and Dmd^mdx-4Cv mouse hearts under unloaded Langendorff conditions which indicated similar GCaMP6f expression and cardiomyocyte arrangement before preload challenge protocol. B: confocal images of subepicardial cardiomyocytes of 3 WT (top) and 3 Dmd^mdx-4Cv (bottom) mouse hearts post-preload challenge protocol. C: summary data of percent GCaMP6f-positive regions in WT (circles) and Dmd^mdx-4Cv (squares) hearts under unloaded Langendorff conditions (combined, n = 3, 7–8-mo WT, and n = 2, 7-mo Dmd^mdx-4Cv) and post-preload challenge. Each symbol is an average of 3–4 sections/heart. **P = 0.004; WT vs. Dmd^mdx-4Cv, one-tailed Mann–Whitney U test via GraphPad Prism, which considers ties among values. n = 5, 3–5-mo WT and n = 5, 4–7-mo Dmd^mdx-4Cv hearts.

Figure 6.

Cellular Ca²⁺ handling abnormalities in dystrophic hearts following sustained preload challenge. Example GCaMP6f Ca²⁺ signals from four representative subepicardial cardiomyocytes of electrically stimulated (3 Hz, dashed line) wild-type (WT; A) and Dmd^mdx-4Cv (B) hearts following sustained preload challenge. Note uniform Ca²⁺ transients in cardiomyocytes of WT (A) and Ca²⁺-wave (cell 1; B) and Ca²⁺-overload (cell 2; B) behavior in Dmd^mdx-4Cv. C: summary data of percent (%) Ca²⁺-overloaded cells in WT (circles) and Dmd^mdx-4Cv (squares) hearts. **P = 0.008; WT vs. Dmd^mdx-4Cv, one-tailed Mann–Whitney U test via GraphPad Prism, which considers ties among values. n = 5, 3–5-mo WT and n = 4, 4–7-mo Dmd^mdx-4Cv hearts. Note only cardiomyocytes with GCaMP6f fluorescence were used to assess %Ca²⁺-overloaded cells.

To investigate arrhythmia incidence in dystrophic hearts following the sustained ventricular preload challenge, an additional cohort of non-GCaMP6f wild-type or dystrophic Dmd^mdx-4Cv hearts were subjected to the preload challenge protocol. Hearts were allowed to beat at sinus rhythm (no electrical stimulation), with cardiac electrical activity monitored using bath ECGs (Fig. 7, A–D). Under unloaded Langendorff conditions, both wild-type and dystrophic hearts had a similar heart rate (Fig. 7E) with a low incidence of ventricular arrhythmias (Fig. 7F). Following preload elevation to 20 mmHg, hearts of both wild-type and dystrophic mice had similar heart rates (Fig. 7G), but hearts of dystrophic mice had an increased incidence of ventricular arrhythmias in the form of PVCs and continuous salvos of PVCs (Fig. 7H).

Figure 7.

Preload-induced ventricular arrhythmias in dystrophic hearts. Example traces of left ventricular (LV) pressure (top) and bath ECG (bottom, P wave and QRS complex marked in 1st cycle) in wild-type (WT, A and B) and Dmd^mdx-4Cv (C and D) hearts. Traces obtained at baseline (0-mmHg preload, A and C) and during the 25–30-min preload challenge (B and D). D: premature ventricular excitation and associated premature ventricular complex (PVC) marked by dashed boxes. Summary data of heart rate (E) and arrhythmia score (F) under baseline conditions (0-mmHg preload) in WT (circles) and Dmd^mdx-4Cv (squares) hearts. Summary data of heart rate (G) and arrhythmia score (H) 25–30 min following 20-mmHg preload challenge. *P = 0.02, WT vs. Dmd^mdx-4Cv, one-tailed Mann–Whitney U test. n = 11, 3–4-mo WT and n = 12, 3–4-mo Dmd^mdx-4Cv hearts.

DISCUSSION

This investigation examined cardiac function and Ca²⁺ homeostasis in a mouse model of muscular dystrophy, which expresses the genetically encoded Ca²⁺ indicator GCaMP6f. An advantage to this experimental approach is that it provides unique insight into Ca²⁺ homeostasis after cardiomyocytes were stretched in their native arrangement within the organ in response to a physiological increase in preload pressure. Importantly, studies were performed in wild-type and dystrophic mice at an age with minimal apparent changes in baseline ex vivo left ventricular function as assessed by cardiac pressure development. These data are consistent with several investigations in various mdx mouse models of muscular dystrophy where overt cardiac dysfunction appears later in the life span (30–33). Expression of the GCaMP6f Ca²⁺ indicator in cardiomyocytes did not appear to affect baseline pressure development as GCaMP6f positive and GCaMP6f negative hearts had a similar left ventricular contractile function in both wild type and dystrophic hearts (Fig. 2). However, there were some observed trends (Fig. 2, F–H and Fig. 3, A–C) that may necessitate further investigation into potential Ca²⁺ buffering of the Ca²⁺ indicator (17, 34, 35). The baseline release of LDH was also similar between GCaMP6f positive and GCaMP6f negative hearts, as well as between wild-type and dystrophic hearts (Fig. 4A). However, using a 30-min preload challenge protocol, significant dysfunction was induced in dystrophic hearts, with increased LDH release (Fig. 4B), loss of cardiomyocyte GCaMP6f fluorescence (Fig. 5), cardiomyocyte Ca²⁺-overload behavior (Fig. 6), and increased incidence of PVCs (Fig. 7). Thus, similar to investigations in other mdx mouse models functional abnormalities can be unmasked at early ages using cardiac stress (36, 37).

Numerous investigations have shown the susceptibility of dystrophic muscle to damage after mechanical stress. Indeed, a standard assay in the skeletal muscle field is a loss in sarcolemmal integrity assessed via entry of an exogenously applied membrane-impermeant substance (e.g., Evans-Blue dye) or loss of endogenously expressed intracellular molecules (e.g., creatine kinase, lactate dehydrogenase) into surrounding tissues and the bloodstream (14). Such assays have also been applied to investigations into cardiac muscle damage although the mechanisms of cardiac damage and resulting cardiomyopathy remain to be clearly resolved (6, 38). Our studies extend this cardiac literature by indicating a significant decrease in cellular GCaMP6f fluorescence in a mouse model with transgenic cytosolic expression of the GCaMP6f Ca²⁺ indicator in cardiomyocytes (Fig. 5), which is consistent with a loss in membrane integrity with mechanical stretch of dystrophin-deficient cardiomyocytes. These findings were complemented by a traditional assay of LDH release into the coronary effluent following sustained preload challenge (Fig. 4). Sarcolemmal disruptions have been shown to play a key role in the excessive entry of extracellular Ca²⁺ into the cell. Our studies show a preload-induced increase in the number cardiomyocytes with Ca²⁺ overload, proarrhythmic Ca²⁺ waves, and Ca²⁺ transients with abnormal kinetics (Fig. 6).

The mechanisms that link muscle stretch, dystrophin deficiency, Ca²⁺ handling abnormalities, and muscle damage continue to be an active area of investigation. The Ca²⁺ influx hypothesis for stretch-induced muscle damage in the setting of dystrophin deficiency (39) remains supported by the field (13, 40, 41). Prominent Ca²⁺ influx pathways in the dystrophic myocardium include membrane microruptures (19, 40), activation of TRPC ion channels (42–45) and TRPV ion channels (10, 46). The excessive Ca²⁺ influx is further exacerbated by stretch-induced ROS production (6, 47) and RyR-mediated SR Ca²⁺ leak (48–50). Thus, although the primary mechanism likely initiates with sarcolemmal dysfunction and unregulated Ca²⁺ influx because of dystrophin deficiency, a vicious feedback cycle begins leading to additional Ca²⁺ influx, aberrant SR Ca²⁺ reuptake or Ca²⁺ release, mitochondrial permeability transition, mitochondrial damage and energy depletion, generation of free radicals and inflammatory mediators, and resulting muscle damage (6, 38, 42).

A predisposition to ventricular arrhythmias in mdx mouse models has previously been reported, with most investigations using programmed electrical stimulation or catecholamine challenge to induce arrhythmias (49, 51). In vivo ECG recordings reveal an increased incidence of PVCs in mdx versus wild-type mice, and following in vivo isoproterenol challenge PVCs transitioned into spontaneous sustained ventricular tachycardia (48). Dystrophin deficiency also associated with alterations in both Ca²⁺ and Na⁺ currents, which may play a role in aberrant electrical activity and/or impulse propagation (52, 53). Under our unloaded Langendorff conditions in ex vivo perfused hearts arrhythmia incidence was low in both wild-type and dystrophic hearts. However, sustained preload alone increased PVC incidence in dystrophic hearts (Fig. 7). Although we did not observe severe ventricular tachycardia or ventricular fibrillation in this investigation, such arrhythmias would likely be triggered by additional exposure to neurohormonal agonists, or in hearts of older mice with fibrosis and/or structural remodeling (41).

Study Limitations and Future Studies

A key limitation of the present investigation is that confocal imaging of cardiomyocyte GCaMP6f fluorescence was performed after, not during, the preload challenge. Thus, the mechanisms of decreased GCaMP6f fluorescence signal of cardiomyocytes and the progression of Ca²⁺ handling dysfunction during the preload challenge remain to be determined. Although GCaMP6f fluorescence can be altered by the intracellular environment [e.g., pH or reactive oxygen species (54)], we have successfully used this indicator during severe ischemia-reperfusion injury (27) and the significant loss of myocyte GCaMP6f signal was unexpected (Fig. 5B). Future experiments examining cardiomyocyte GCaMP6f fluorescence in real time will provide additional insight into if GCaMP6f signal declines slowly consistent with minor disruptions in membrane integrity, or alternatively, if progressive Ca²⁺ overload leads to cardiomyocyte death and a rapid decline in GCaMP6f fluorescence because of catastrophic loss of membrane integrity. In addition, real-time measurements of Ca²⁺ handling may provide insight on the disconnect between the loss of GCaMP6f fluorescence in the subepicardium of dystrophic hearts (Fig. 5) yet with preserved contractile function during the sustained preload challenge (Fig. 3). Although speculative, it is possible that during the preload challenge populations of cardiomyocytes exhibit elevated Ca²⁺ transients and hypercontractility, which may counteract cardiac damage observed in other cardiomyocyte regions (Figs. 4 and 5) to preserve overall contractile function. Single-photon confocal imaging was used to assess GCaMP6f fluorescence, and this methodology does not permit imaging of cardiomyocytes deeper within the heart tissue. Additional studies using multiphoton fluorescence microscopy (29) will allow for the investigation of cardiomyocyte structure and function in the midmyocardium and will determine if loss of cardiomyocyte GCaMP6f and Ca²⁺ handling abnormalities are present throughout the left ventricular wall. Furthermore, the right ventricle is thin compared with the left ventricle, and right ventricular remodeling precedes left ventricular remodeling in the mdx model (33, 55). Thus, the right ventricle may be more susceptible to preload-induced cardiomyocyte dysfunction. Future investigations will probe key differences in physiology and pathophysiology following stretch between the respective chambers.

Conclusions

With the use of cardiomyocyte-specific expression of the GCaMP6f Ca²⁺ indicator in mice of a young age, this investigation provides additional support for the stretch-induced Ca²⁺ damage hypothesis (13, 40). Although the present investigation did not identify mechanisms of Ca²⁺ overload induced by acute ventricular preload elevation, the methodology developed herein will serve as an ideal experimental platform to test preload-induced alterations in cardiomyocyte Ca²⁺ flux balance. Such investigations will accelerate therapeutics to restore sarcolemmal function or pharmacologically reduce Ca²⁺ stress on the cardiomyocyte to prevent pathological Ca²⁺ handling, adverse ventricular remodeling, and arrhythmia in DMD.

DATA AVAILABILITY

The data underlying this article will be shared on reasonable request to the corresponding author.

GRANTS

This work was supported by the National Institutes of Health Grants R01HL136292 (to T.L.D.) and R01AR070517 (to D.D.), United States Department of Defense Grant MD210064 (to D.D.), and University of Missouri System Tier 2 Award, Research and Creative Works Strategic Investment Program (M.K. and K.S.M., coprincipal investigators).

DISCLOSURES

D.D. is a member of the scientific advisory board for Solid Biosciences and an equity holder of Solid Biosciences. D.D. is a member of the scientific advisory board for Sardocor Corp. The Duan laboratory received research support unrelated to this project from Solid Biosciences in the last 3 years. The Duan laboratory has received research support unrelated to this project from Edgewise Therapeutics in the last 3 years. None of the other authors has any conflicts of interest, financial or otherwise, to disclose.

AUTHOR CONTRIBUTIONS

V.H., Z.N., M.D.L., L.M.H., M.K., C.P.B., D.D., K.S.M., and T.L.D. conceived and designed research; V.H., Z.N., M.D.L. M.K., and T.L.D. performed experiments; V.H., Z.N., E.M.B., M.D.L., M.K., and T.L.D. analyzed data; V.H., Z.N., E.M.B., M.D.L., L.M.H., M.K., C.P.B., D.D., K.S.M., and T.L.D. interpreted results of experiments; V.H., Z.N., E.M.B., and T.L.D. prepared figures; V.H., Z.N., and T.L.D. drafted manuscript; V.H., Z.N., E.M.B., L.M.H., M.K., C.P.B., D.D., K.S.M., and T.L.D. edited and revised manuscript; V.H., Z.N., E.M.B., M.D.L., L.M.H., M.K., C.P.B., D.D., K.S.M., and T.L.D. approved final version of manuscript.