Cardiac myopathies are the second leading cause of death in patients with Duchenne and Becker muscular dystrophy, the two most common and severe forms of a disabling striated muscle disease. Although the genetic defect has been identified as mutations of the dystrophin gene, very little is known about the molecular and cellular events leading to progressive cardiac muscle damage. Dystrophin is a protein linking the cytoskeleton to a complex of transmembrane proteins that interact with the extracellular matrix. The fragility of the cell membrane resulting from the lack of dystrophin is thought to cause an excessive susceptibility to mechanical stress. Here, we examined cellular mechanisms linking the initial membrane damage to the dysfunction of dystrophic heart.
Cardiac ventricular myocytes were enzymatically isolated from 5- to 9-month-old dystrophic mdx and wild-type (WT) mice. Cells were exposed to mechanical stress, applied as osmotic shock. Stress-induced cytosolic and mitochondrial Ca2+ signals, production of reactive oxygen species (ROS), and mitochondrial membrane potential were monitored with confocal microscopy and fluorescent indicators. Pharmacological tools were used to scavenge ROS and to identify their possible sources. Osmotic shock triggered excessive cytosolic Ca2+ signals, often lasting for several minutes, in 82% of mdx cells. In contrast, only 47% of the WT cardiomyocytes responded with transient and moderate intracellular Ca2+ signals. On average, the reaction was 6-fold larger in mdx cells. Removal of extracellular Ca2+ abolished these responses, implicating Ca2+ influx as a trigger for abnormal Ca2+ signalling. Our further experiments revealed that osmotic stress in mdx cells produced an increase in ROS production and mitochondrial Ca2+ overload. The latter was followed by collapse of the mitochondrial membrane potential, an early sign of cell death.
Overall, our findings reveal that excessive intracellular Ca2+ signals and ROS generation link the initial sarcolemmal injury to mitochondrial dysfunctions. The latter possibly contribute to the loss of functional cardiac myocytes and heart failure in dystrophy. Understanding the sequence of events of dystrophic cell damage and the deleterious amplification systems involved, including several positive feed-back loops, may allow for a rational development of novel therapeutic strategies.
Dystrophinopathies result from mutations of the dystrophin gene (reviewed in Ervasti and Campbell1). Dystrophin is a 427 kDa protein linking the cytoskeleton to a complex of transmembrane proteins, which interact with the extracellular matrix. Duchenne and Becker muscular dystrophy (DMD and BMD) are the two most common and severe forms and are characterized by progressive and disabling muscle weakness (one case per 3500 boys). Cardiac manifestations are present in essentially all patients by 20 years of age. The most common abnormality is dilated cardiomyopathy. Around 20% of the patients succumb from the cardiac manifestations. Clinically, the disease initially shows ECG abnormalities. Later, echocardiographic examinations reveal ventricular wall motion abnormalities in regions of fibrosis. With spreading of fibrosis, ventricular dysfunction and arrhythmias occur, ultimately leading to heart failure and sudden death.2 With improving medical care for the skeletal muscle weakness, the cardiac disease becomes more and more limiting for the survival of the patients.
In the skeletal muscle field, there is a significant body of data describing the cellular pathophysiological events (for review see Allen et al.3). It is generally believed that the dystrophin deficiency leads to exaggerated sarcolemmal fragility. Abnormal Ca2+ influx and intracellular Ca2+ homeostasis after mechanical stress have been implicated in progressive death of dystrophic skeletal muscle fibres.4,5 The mdx mouse is a well-established model for skeletal muscle dystrophy, and it also develops a cardiac phenotype with age that is similar to the alterations observed in humans during dystrophic cardiomyopathy.6 It is suggested that in skeletal muscle Ca2+ entry could occur via voltage-independent pathways,3 such as ‘leak’ channels,7 stretch-activated channels (SAC),8 or store operated channels (SOC).9 The molecular identity of these channels is not yet fully clear, but some of them are thought to contain TRP channel subunits.3 In addition, there are findings suggesting that the vulnerability of mdx muscle leads to microruptures which may also allow excessive Ca2+ influx.10 Despite all these identified possible pathways for Ca2+ entry, the main question still remains open: what is the sequence of cellular events that initiate Ca2+ influx and which are the positive feed-back loops that amplify this influx and lead to Ca2+ overload and cell death?
Identical questions apply to the cardiac phenotype, but much fewer studies were carried out to understand cellular mechanisms leading to cardiac pathology. However, the clinical findings described above and some experimental studies on the organ level also indicate an abnormal sensitivity of cardiomyocytes to mechanical stress, for example, after aortic banding11 or prolonged exposure to β-adrenergic stimulation to induce arterial hypertension.12 In addition, histological examination revealed replacement of muscle with connective tissue and the presence of apoptotic and necrotic myocytes, also in cardiac muscle.13,14 Thus, cell death most likely underlies the development of fibrosis and progressive deterioration of both tissues.
A functional study found a high stress sensitivity of isolated mdx cardiomyocytes and that the cell damage could be suppressed by the membrane protective detergent Poloxamer P188.15 In addition, a report suggested SAC to contribute to the abnormal Ca2+ influx pathway in mdx myocytes.16 Here, we used confocal imaging with various fluorescent indicators to monitor intracellular processes that may link the initial transmembrane Ca2+ influx to cell damage. We observed that mdx cardiac myocytes exhibit an extreme Ca2+ signalling response to mechanical stress. More importantly, we identified an abnormal activation of the Ca2+-induced Ca2+ release signalling pathway together with an increased generation of reactive oxygen species (ROS) as two synergistic mechanisms contributing to the amplification of these Ca2+ signals. Preliminary results of these studies have been published as an abstract.17
Ventricular myocytes were enzymatically isolated from 5- to 9-month-old mice as previously described.18,19 The study conforms with the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996). C57BL/10ScSn-mdx and wild-type (WT) mice with identical genetic background (5–10 months old) were provided by Drs M. Rüegg (University of Basel, Switzerland), U. Rüegg (University of Geneva, Switzerland), or purchased from the Jackson laboratory (USA).
The isotonic superfusion solution contained (in mM): 140 NaCl, 5.4 KCl, 1.8 CaCl2, 1.1 MgCl2, 5 HEPES, and 10 glucose. The osmolality was ∼310 mosm/kg, and pH 7.3. The hypotonic solution contained 70 mM NaCl instead of 140 mM, had osmolality ∼170 mosm. Zero Ca2+ solution contained 2 mM EGTA. Where indicated, apocynin or ryanodine and thapsigargin were added for at least 30 min before the experiment. Most chemicals were obtained from Sigma. Mn-cpx 3 and apocynin were from Calbiochem, ryanodine from Alamone. All experiments were performed at room temperature (20–22°C).
Changes in cytosolic and mitochondrial [Ca2+], ROS production, and mitochondrial membrane potential were monitored with fluorescent indicators fluo-3 AM (5 µM), rhod-2 AM (5 µM), fura-red AM (5 µM), 5-(and-6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CM-H2DCFDA; 20 µM), and tetramethylrhodamine ethyl ester (TMRE; 100 nM), as previously described.20–22 As a rule, cells were incubated with indicators for 30 min at room temperature, followed by at least 15 min for de-esterification. Resting [Ca2+] and Ca2+ influx were measured with a double-indicator ratiometric procedure, which simultaneously utilizes fluo-3 and fura-red indicators to correct for dye dilution during osmotic swelling.23 Fluo-3 AM was from Biotium, other indicators were from Invitrogen. A laser-scanning confocal microscope (Radiance, Bio-Rad) was used to acquire confocal images.22,24 Fluo-3, fura-red, and CM-H2DCFDA were excited with the 488 nm line of an Argon laser. TMRE and rhod-2 were excited with a HeNe laser at 543 nm. The emitted light was collected above 500 nm and above 570 nm, respectively. The emission of fluo-3 and fura-red was simultaneously recorded with two PMTs using 515/30 and >600 nm emission filters and the ratio calibrated according to.23
Analysed data are represented as mean ± SEM. Where stated, statistical significance was determined using Student's t-test or the log-rank test. In the figures, single asterisk indicates P < 0.05, double asterisks indicate P < 0.01.
Osmotic shock triggers excessive cytosolic Ca2+ signals in mdx but not wild-type cardiomyocytes
We applied osmotic shocks to mimic some of the pathophysiological conditions of mechanical stress encountered by the cell in vivo.5,25 Hypotonic solution induces cell swelling and mechanical deformation of the cell membrane. Cytosolic Ca2+ transients were recorded in both WT and mdx cardiomyocytes loaded with fluo-3. As a rule, series of 100 XY confocal scans were acquired at 0.5 Hz. Exposure of the cardiac myocytes to hypotonic solution (170 mosm) for 40 s consistently resulted in reversible cell swelling (by 15.8 ± 1.5%, n = 34). In 82% of mdx myocytes (19 out of 23 cells), the return to isotonic solution caused extreme cytosolic Ca2+ signals such as bursts of Ca2+ sparks and repetitive Ca2+ waves exhibiting amplitudes comparable with normal Ca2+ transients (as shown in Figure 1A, left panel). In the majority of mdx cells, these Ca2+ signals lasted for the full duration of the recording (up to 3 min). In contrast, in only 47% of the WT cardiomyocytes, osmotic shock produced Ca2+ signals (nine out of 19 cells) and these were mostly moderate and short-lived (Figure 1A, right panel). The typical Ca2+ responses in mdx and WT cells are also visualized in movies in the online supplement. Figure 1B shows the averaged changes in intracellular fluorescence in all studied WT and mdx cardiomyocytes, normalized to the resting values before osmotic shock. The stress-induced increase in fluorescence was larger and longer lasting in mdx cells compared with WT myocytes. The mean cellular fluorescence signal recorded during the first 60 s after returning to the iso-osmotic solution was 6-fold larger in mdx cells (Figure 1C).
A small Ca2+ influx is required for stress-induced Ca2+ signals in mdx cardiomyocytes
The above observation together with the recent results by Yasuda et al.15 suggest that mdx myocytes are more susceptible to membrane damage than WT cardiomyocytes. To test whether these Ca2+ signals depend on extracellular Ca2+, we briefly removed Ca2+ from the extracellular solution. Figure 2 shows that switching to a solution containing 0 mM Ca2+ completely abolished the response to osmotic shock. This observation confirms that Ca2+ entry is required, although the precise pathway(s) leading to Ca2+ influx into dystrophic cells are yet to be established and seems to involve multiple molecular entities.3
Nevertheless, the increase in plasmalemmal permeability to Ca2+ and the slightly elevated resting Ca2+ concentration in mdx myocytes (see Figure 2E and Williams and Allen16) alone are unlikely to explain the sustained Ca2+ signals in mdx cells subjected to osmotic shock. In cells treated with ryanodine (5 µM) and thapsigargin (500 nM) to eliminate the function of the sarcoplasmic reticulum (SR), a substantial Ca2+ influx during the superfusion with hypotonic solution would be visible as an increase of the cytosolic Ca2+ concentration. However, when applying a ratiometric indicator technique involving fluo-3 and fura-red to compensate for dye dilution due to osmotic swelling, only a modest elevation of the Ca2+ concentration was observed, suggesting that the Ca2+ influx was small (Figure 2C and D, gray circles).
Osmotic shock stimulates reactive oxygen species production
Because we were not able to detect a substantial Ca2+ influx during the shock, we suspected some intracellular mechanisms by which even small increases in [Ca2+]i due to Ca2+ influx were amplified. One possibility could be oxidation of the cytosolic environment due to excessive ROS production,26 which could affect several proteins involved in intracellular Ca2+ homeostasis, most notably the ryanodine receptors (RyRs), thereby enhancing the sensitivity of SR Ca2+ release.
To test whether osmotic shock enhances ROS generation, we loaded myocytes with CM-H2DCFDA. In cells, this non-fluorescent compound is hydrolysed to DCFH and ROS generation is detected as a result of DCFH oxidation to DCF. Figure 3A shows images from one mdx cardiac myocyte before (image at 20 s), during (50 s), and after osmotic shock (80, 110, 140, and 170 s). Insets illustrate the banded fluorescence typical of a mitochondrial distribution. Figure 3B represents averaged and normalized changes in DCF fluorescence in mdx and WT cells. To quantify the stress-induced changes in ROS production, we performed a linear fit to the fluorescence signal in each cell before and after osmotic shock. The slope of the DCF signal reflects the rate of ROS production. Before the osmotic shock, the ROS-related signals rose monotonically, presumably due to a basal generation of ROS. The initial slope was significantly larger in mdx myocytes than in WT cells (0.70 ± 0.09, n = 12 and 0.40 ± 0.07, n = 9, respectively) indicating an oxidative stress in these cells, as also suggested in Williams and Allen.27 Furthermore, we observed that the slope increased after the shock in both WT and mdx cells, but more in mdx myocytes than in WT cells [ratio of slopes before to after the shock was 1.72 ± 0.2 (n = 12) and 1.34 ± 0.12 (n = 9) in mdx and WT, respectively].
To confirm the involvement of ROS-dependent mechanisms in the generation of stress-induced Ca2+ signals, we incubated mdx myocytes with 10 µM Mn-cpx3, a superoxide dismutase mimetic. Figure 4A shows that this ROS scavenger nearly eliminated cytosolic Ca2+ transients produced by osmotic shock. After washing out the scavenger for 30 min, five out of six mdx cardiomyocytes recovered their exacerbated response. Figure 4B shows a significant decrease in the mean intracellular fluorescence recorded during the first 60 s after returning to an iso-osmotic solution in the presence of the scavenger.
There are several possible sources of ROS in cardiac myocytes that may be activated in response to mechanical stress. NAD(P)H oxidase (NOX) is known to be overexpressed in dystrophic heart27 and was found to be rapidly activated by osmotic swelling in astrocytes.28 Therefore, we tested whether NOX could be a source for a rapid increase in the ROS production in response to the mechanical stress in mdx myocytes. Cells were incubated for 30 min with the NOX inhibitor apocynin (0.2–1 mM). Figure 4C and D shows that the drug nearly eliminated the intracellular Ca2+ signals induced by osmotic shock, suggesting NOX as a possible source of acute ROS production.
Excessive cytosolic Ca2+ is taken up by the mitochondria
Mitochondria are well-established sources of ROS in myocardium subjected to acute or chronic mechanical stress.29 Mitochondrial ROS could be responsible for the sustained cellular Ca2+ signals observed in mdx myocytes (note the gradual increase in ROS-related fluorescence in the mitochondrial regions later during the experiment shown in Figure 3). It is known that muscle cells continuously generate ROS, as a by-product of the mitochondrial respiratory chain, and that enhanced mitochondrial Ca2+ uptake stimulates further mitochondrial ROS production. We tested whether and to what extent mitochondria take up cytosolic Ca2+ after osmotic shock.
Mdx and WT myocytes were loaded with the Ca2+ indicator rhod-2 and imaged with the identical experimental protocol. Rhod-2 is a charged molecule that preferentially partitions into the mitochondria. Figure 5A shows the mitochondrial rhod-2 distribution and the fluorescence signal in mdx myocytes before (image at 1 s), during (image at 28 s), and after the shock. Following 40 s of osmotic shock, fluorescence gradually increased within all mitochondria. Figure 5B shows the time course of rhod-2 signals in three groups of mitochondria indicated in A. In some mitochondria, the rhod-2 signal, after an initial rise, decreased at later times, indicating a loss of rhod-2 and Ca2+. Figure 5C represents averaged normalized changes in rhod-2 fluorescence in six mdx and eight WT cells, indicating that following osmotic stress mitochondria in mdx myocytes sequester significantly more Ca2+ which can add to the more pronounced ROS production.
Osmotic shock induces mitochondrial depolarization in mdx cardiac myocytes
It is established that mitochondrial Ca2+ overload and the subsequent oxidative stress can lead to collapse of the mitochondrial membrane potential (Δψm) and/or to the opening of the mitochondrial permeability transition pore (mPTP), ultimately causing necrotic/apoptotic cell death.29 Our results indicate that mitochondrial Ca2+ uptake is enhanced in mdx cardiac myocytes. We also found that some mitochondria lose Ca2+ after a period of initial accumulation, possibly as a result of irreversible mitochondrial depolarization.30 Therefore, we carried out experiments to directly monitor changes in mitochondrial membrane potential. TMRE is a charged dye, which distributes across the mitochondrial membrane in a voltage-dependent manner. Whenever the potential across the mitochondrial membrane dissipates, TMRE fluorescence is lost. Figure 6A shows one mdx cardiomocyte loaded with 100 nM TMRE. A single mitochondrion (or a small group of mitochondria) can be visualized in each of the marked boxes. Subsequent to the osmotic shock, some mitochondria displayed irreversible loss of Δψm whereas others did not. Changes in TMRE fluorescence for four individual mitochondria are plotted in Figure 6B. Note that the final level of TMRE signal is essentially the same in all four organelles, indicating their complete depolarization. Figure 6C compares the incidence and timing of osmotic shock-induced mitochondrial depolarization in mdx and WT cardiomyocytes. For each time point, we calculated the number of polarized mitochondria in all cells of each type and normalized it to the total numbers of organelles imaged. In mdx cells, osmotic shock induced depolarization of ∼2% of the mitochondria within 400 s of experimental recording (black line). Loss of TMRE signals by individual mitochondria was observed already 4 s after the shock. In contrast, in WT myocytes, the fraction of depolarized mitochondria was much smaller (0.3%) and the onset was later (100 s after the shock). It should be mentioned that in some mdx myocytes shock induced severe mitochondrial depolarization waves that resulted in complete collapse of Δψm across the entire cell within a few seconds. These cells were not included in the analysis presented in Figure 6C.
Dystrophin-deficient cardiomyocytes are vulnerable to mechanical stress. Recently published work connects increased membrane fragility and stretch-induced Ca2+ influx with death of dystrophic cells.15 In the present study, we identified two cellular signal amplification mechanisms that may link the fragility of the mdx cardiomyocyte membrane to later cell damage. As detailed below, the simultaneous activation of these two signal amplification systems is detrimental, because of bidirectional cross-talk with positive feed-back. Thus, the cellular response proceeds beyond the physiological limits, culminating in a chain of events, which may later end in apoptotic and/or necrotic cell death.
The massive Ca2+ signals observed in mdx myocytes after osmotic shock were manifest as a surge of abundant spontaneous Ca2+ sparks, regularly blending into Ca2+ waves travelling along the cell. In resting cardiomyocytes such signals are the hallmark of abnormal Ca2+-induced Ca2+ release (CICR) activation and are commonly initiated by overloading the SR with Ca2+ in the course of various noxious conditions. The importance of CICR for the observed signal amplification was also emphasized by the absence of such Ca2+ signals in cells pretreated with ryanodine and thapsigargin. However, for Ca2+ overload to develop in a short period of time (i.e. during the exposure to hypotonic solution), Ca2+ influx has to be substantial. Surprisingly, such a massive Ca2+ influx could not be detected in our experiments, even though the behaviour of the myocytes in zero Ca2+ indicated an important role for at least some minor transsarcolemmal Ca2+ influx. The obvious discrepancy between massive and rapid CICR activation despite minimal Ca2+ entry and only slightly elevated resting Ca2+ levels led us to consider additional mechanisms that could make the RyRs rapidly more sensitive and provoke spontaneous Ca2+ signals without pronounced SR Ca2+ overload.
ROS have been implied in changing gating and Ca2+ sensitivity of the RyRs via redox modifications of the channels.31 Furthermore, increased ROS generation after mechanical stretch has been reported even in normal cardiac muscle.26 Our findings revealed an elevated basal rate of ROS production in mdx cells. Furthermore, mechanical stress led to enhanced ROS production in all cardiomyocytes, but significantly more pronounced in mdx cells. Although the first finding may underlie the propensity of mdx cells to overreact after small Ca2+ influx signals, the latter may be a consequence of the larger Ca2+ signals in these cells. As one might expect based on these findings, ROS scavengers reversibly suppressed the excessive stress-induced Ca2+ signalling in mdx cells. Taken together, our results suggest that the change of the cytosolic oxidative environment due to increased [ROS] promotes the abnormal CICR activation without pronounced Ca2+ overload.
ROS generation in response to mechanical forces may originate from several sources, including the NOX, xanthine oxidase, or other oxidase systems, such as nitric oxide oxidase, and mitochondria. Our results with the NOX inhibitor apocynin suggest an involvement of this source of ROS. Owing to its overexpression in dystrophic heart27 and due to its rapid activation by osmotic swelling,28 this source of ROS may be particularly important in the early phases of the response. It is also established that an elevation in mitochondrial [Ca2+] subsequent to prolonged cytosolic Ca2+ signal activity stimulates mitochondrial ROS production.29 Consistent with this notion, we observed increased Ca2+ levels in cardiac mitochondria following the osmotic shock that were significantly larger in mdx cells.
Cross-talk between Ca2+-induced Ca2+ release and reactive oxygen species
CICR and ROS generation are two signalling pathways that exhibit a high degree of positive feed-back. Either mechanism could lead to the observed massive Ca2+ signals. However, the combination of both pathways being activated together may have even more severe consequences, as both pathways exhibit mutual and synergistic cross-talk with each other. For example, ROS could promote RyRs openings as well as further increase membrane fragility due to lipid peroxidation. In addition, ROS could stimulate Ca2+ influx into the cells via SAC and/or SOC, as some TRP channels (potential candidates for SOC and SAC) have been shown to be sensitive to the redox potential of the cytosol.3 On the other hand, CICR and elevated [Ca2+]i favour mitochondrial ROS production, as discussed above. Taken together, the simultaneous activation of these signalling pathways is particularly destructive for the cells because it has a tendency to escalate into mutually synergistic positive feed-back signalling loops.
Link between abnormal cellular signalling and pathology of the heart
Although WT myocytes survive the initial Ca2+ overload, the strong activation of several pathways may overwhelm the cellular defence mechanisms in mdx cells. It has been postulated for many years that proteases are a downstream Ca2+-activated target involved in protein degradation and necrosis of dystrophic skeletal muscle.3 Calpain is considered to be a likely candidate, as its expression is greater in mdx muscle.3 Because calpain is expressed in heart, and because calpain inhibitors protect from ischaemia–reperfusion injury,32 it is likely that this protease forms a part of the cellular death pathway in mdx heart.
Under pathological conditions, cytosolic and mitochondrial Ca2+ overload and/or excess in ROS can lead to opening of the mPTP, which is accompanied by a dissipation of the mitochondrial membrane potential and usually precedes necrotic/apoptotic cell death.29,33 Our data reveal multiple mitochondrial dysfunctions in mdx cardiac myocytes following osmotic stress. In particular, they show mitochondrial Ca2+ overload, increased ROS production, irreversible depolarization of the mitochondrial membrane, which can be a direct indication of mPTP opening. Apoptotic and necrotic cells have been observed in cardiac tissue of dystrophy patients and mdx mice. Lost cardiomyocytes cannot regenerate and are likely to be replaced by connective tissue, ultimately leading to the observed fibrosis.2
At present, several studies are conducted to develop treatments of the cardiac dystrophic phenotype by either replacing the lost cardiomyocytes with stem cell approaches34 or to make the myocytes mechanically less vulnerable by gene transfer of dystrophin, utrophin, or functional dystrophin fragments (microdystrophins).35 Notably, mdx mice can partly compensate for the lack of dystrophin by adaptive upregulation and redistribution of the protein utrophin.36 An mdx mouse lacking utrophin presents with a more severe phenotype,37 particularly in skeletal muscle, but also in cardiac muscle.38 Although cell replacement therapies will obviously only be able to treat late stages of the disease, curative gene therapy may require a long time to develop for routine application. In the meantime, it would be helpful to implement pharmacological treatment options which are able to slow down the progression of the disease.4 Considering the early cellular stress responses reported here, this might be accomplished by interfering with any of the early cellular events leading to excessive Ca2+ and ROS signals.
Supplementary material is available at Cardiovascular Research online.
Swiss Foundation for Research on Muscle Diseases; Muscular Dystrophy Association; Swiss National Science Foundation; Swiss State Secretariat for Education and Research; UMDNJ & Sigrist Foundations.
Conflict of interest: none declared.
We thank Drs Mohammed Fanchaouy, Larry Gaspers, Jakob Ogrodnik, Roman Shirokov, and Andrew Thomas for discussions and Daniel Lüthi for technical assistance.
- cardiac myocyte
- signal transduction
- extracellular matrix
- becker's muscular dystrophy
- heart failure
- cause of death
- cell death
- cell membrane
- tissue membrane
- microscopy, confocal
- precipitating factors
- reactive oxygen species
- stress, mechanical
- cell injury
- integral membrane proteins
- muscle cells
- ventricular myocyte
- gene abnormality
- mitochondrial membranes
- dystrophin gene
- muscle, striated