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Human Molecular Genetics Advance Access originally published online on May 25, 2005
Human Molecular Genetics 2005 14(14):1921-1933; doi:10.1093/hmg/ddi197
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© The Author 2005. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oupjournals.org

Cardiomyopathy in dystrophin-deficient hearts is prevented by expression of a neuronal nitric oxide synthase transgene in the myocardium

Michelle Wehling-Henricks1, Maria C. Jordan3, Kenneth P. Roos3, Bo Deng4 and James G. Tidball1,2,4,*

1Department of Physiological Science, 2Department of Pathology and Laboratory Medicine, 3Department of Physiology and 4Molecular, Cellular and Integrative Physiology Program, David Geffen School of Medicine, 5833 Life Science Building, University of California, Los Angeles, CA 90095, USA

* To whom correspondence should be addressed. Tel: +1 3102063395; Fax: +1 3108258489; Email: jtidball{at}physci.ucla.edu

Received January 26, 2005; Revised April 18, 2005; Accepted May 17, 2005


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Null mutation of dystrophin causes the lethal pathology of Duchenne muscular dystrophy (DMD) in which there is progressive pathology of skeletal and cardiac muscles. A large proportion of DMD patient deaths are attributable to cardiac dysfunction associated with ventricular fibrosis, arrhythmias and conduction abnormalities, although the relationships between the dystrophin mutation and the cardiac defects are unknown. Here, we tested whether cardiac pathology in dystrophin-deficient mdx mice can be corrected by the elevated production of nitric oxide (NO) by the myocardium. Dystrophin-deficient mdx mice were produced in which there was myocardial expression of a neuronal nitric oxide synthase (nNOS) transgene. Expression of the transgene prevented the progressive ventricular fibrosis of mdx mice and greatly reduced myocarditis. Electrocardiographs (ECG) attained by radiotelemetry of freely ambulatory mice showed that mdx mice displayed cardiac abnormalities that are characteristic of DMD patients, including deep Q-waves, diminished S:R ratios, polyphasic R-waves and frequent premature ventricular contractions. All of these ECG abnormalities in mdx mice were improved or corrected by nNOS transgene expression. In addition, defects in mdx cardiac autonomic function, which were reflected by decreased heart rate variability, were significantly reduced by nNOS transgene expression. These findings indicate that increasing NO production by dystrophic hearts may have therapeutic value.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Cardiomyopathy frequently causes mortality in Duchenne muscular dystrophy (DMD) patients. More than 30% of DMD deaths reported between 1985 and 1989 resulted from cardiac failure (1Go). This figure is surely increasing as previously fatal respiratory deficits are treated with ventilatory assist devices (2Go) permitting cardiac involvement to emerge as a more prevalent feature of the disease. Current management options for patients with dystrophin-deficient cardiomyopathy are typically limited to the use of drugs prescribed to relieve symptoms of congestive heart failure (3Go,4Go). Unfortunately, DMD patients are largely inactive due to loss of ambulation in their early teens and the increasing severity of the cardiomyopathy can undergo unnoticed, untreated and result in sudden death. The dystrophic cardiomyopathy also affects female carriers of dystrophin mutations who are generally spared from the severe skeletal muscle pathology. More than 50% of female carriers experience clinically relevant cardiac involvement often severe enough to merit heart transplantation (5Go,6Go,7Go).

The DMD cardiomyopathy presents clinically and histologically with characteristic features. Electrocardiography (ECG) of DMD patients shows tall right precordial R-waves, decreased S:R ratios, deep Q-waves in the limb and left precordial leads, and often, polyphasic R-waves (8Go–12Go). As the cardiomyopathy progresses, arrhythmias, conduction abnormalities, regional left ventricular wall motion abnormalities, left ventricular dilation and cardiac autonomic dysfunction occur (8Go–10Go,13Go–20Go). Preclinical abnormalities are detectable by ECG in 59% of patients between 6 and 10 years of age and progressively develop into clinically apparent cardiomyopathy in 100% of patients over 18 years of age (21Go). Postmortem examination of cardiac tissue from DMD patients shows that extreme fibrosis, generally localized to the posterobasal wall of the left ventricle, is the most prevalent histological feature of the pathology, while fibrosis of the peripheral conduction system has also been observed (1Go,8Go–10Go,13Go,15Go). Furthermore, remaining viable cardiomyocytes can become encased in connective tissue, thereby compromising their intercellular connections and ability to conduct signals (22Go). The electrical and functional cardiac abnormalities in DMD patients have been attributed to cardiac fibrosis, which suggests that fibrosis is an important component of cardiomyopathy in DMD (8Go).

Hearts of dystrophin-deficient mdx mice share many features of the DMD cardiomyopathy. Similar to DMD patients, mdx mice experience a progressive development of cardiac defects, although the pathology is less severe. Young, adult mice show little pathology at 2–3 months of age, while function and morphology markedly deteriorate by 12 months of age (23Go–25Go). Like DMD patients, mdx mice display ECG abnormalities, autonomic dysfunction, impaired conduction, arrhythmias, deteriorated left ventricular function and dilated cardiomyopathy (23Go,24Go,26Go,27Go). Mdx hearts also experience a severe, progressive accumulation of connective tissue (27Go,28Go), suggesting that fibrosis may also be responsible for some features of the mdx cardiomyopathy.

Pathological fibrosis is also a prominent feature in dystrophin-deficient skeletal muscle (28Go–30Go) and may indicate that common mechanisms underlie fibrosis of dystrophin-deficient hearts and muscles. Because dystrophin-deficient skeletal muscles and hearts experience inflammation (27Go,31Go–36Go), and inflammatory cells are known to promote fibrosis in numerous pathologies (37Go–39Go), it is possible that fibrosis of dystrophic tissues results, in part, from inflammatory processes. This possibility has been supported by the finding that mdx mice rendered T-cell deficient by breeding onto the nu/nu background showed less muscle and cardiac fibrosis than mdx mice with T-cells (28Go). Inflammatory cells may induce fibrosis by secreting cytokines such as transforming growth factor-ß (TGF-ß), which stimulate connective tissue production by fibroblasts (40Go,41Go). Previous investigators have shown that TGF-ß is elevated in the skeletal muscles of dystrophin-deficient mice, dogs and humans (42Go–44Go) during the stages of muscle pathology in which fibrosis occurs and that TGF-ß blockade can reduce collagen expression in mdx diaphragms (40Go).

If cardiac inflammation contributes significantly to myocardial fibrosis and subsequent functional defects in dystrophin-deficient hearts, then interventions that affect the inflammatory process could be valuable in reducing cardiomyopathy in DMD and mdx dystrophy. In our previous work, we showed that inflammation of dystrophin-deficient skeletal muscle is exacerbated by the loss of nNOS from muscle (33Go), which occurs as a secondary consequence of dystrophin-deficiency (45Go,46Go). Expression of a nNOS transgene in mdx mice skeletal muscle to normalize the levels of muscle (nitric oxide) NO production caused great reductions in muscle inflammation and in muscle membrane lysis (33Go), which indicated that muscle-derived NO can serve anti-inflammatory and cytoprotective roles in dystrophic skeletal muscle.

Previous investigators have shown that the hearts of mdx mice lose ~80% of normal nNOS activity (23Go), which is similar to the loss of NOS activity that occurs in mdx skeletal muscle (46Go). This similarity suggests that nNOS defects may also contribute to cardiomyopathy in dystrophin-deficiency. However, there are important distinctions between the distribution and the molecular associations of nNOS in cardiac and skeletal muscles, which can affect the functions of NO in the two muscle types. For example, nNOS in skeletal muscle appears to be localized primarily at the sarcolemma through its association with the dystrophin-complex (45Go). In cardiac muscle, nNOS is located at the sarcolemma (47Go) but has also been reported to be localized at the sarcoplasmic reticulum (48Go) and mitochondria (49Go,50Go), where it is not co-distributed with dystrophin. Thus, the deficiency in nNOS activity in cardiac muscle may primarily reflect defects in its regulation rather than loss of binding to the dystrophin-complex. Predicting the effects of perturbed NOS activity in the heart is more complex because the specific site of NO production in cardiac muscle may have antagonistic effects on the functions mediated by NO. Although NO generated by endothelial NOS at the sarcolemma can have a negative inotrophic effect in cardiac muscle, NO generated by nNOS at the sarcoplasmic reticulum membrane can have a positive effect on contractility (51Go). Competing functions of NO-derived from different NOS isoforms are not known to occur in skeletal muscle.

In the present investigation, we test whether modifications in NO production by nNOS in cardiomyocytes can affect the pathology of mdx hearts. In our hypothetical model, experimental elevation of myocardial NO production in dystrophin-deficient hearts would reduce cardiac inflammation, decrease fibrosis and, as a consequence, normalize those features of cardiac function that have been attributed to fibrosis of the mdx or DMD myocardium. We test our hypothesis by generating a dystrophin-deficient mice line in which there is myocardial expression of a nNOS transgene. The effect of elevated myocardial NO production by nNOS in mdx hearts is assessed by telemetric monitoring of cardiac function in freely ambulatory mice and by testing whether increases in cardiac inflammation and fibrosis are affected.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
nNOS transgene insertion increases nNOS expression and activity in cardiac tissue
Western blot analysis of hearts from wild-type C57 mice and mdx mice at 3, 5, 9 or 18 months of age showed that there is no difference in nNOS concentration between wild-type and mdx hearts at any age that was assayed (Fig. 1A shows data from 5-month-old mice). The concentration of nNOS in the hearts of nNOS transgenic, dystrophin-expressing mice (nNOS Tg+ mice) was 9.4 times the concentration in the hearts of non-transgenic littermates (nNOS Tg– mice) (Fig. 1B) and 2.0-fold greater than the expression levels in nNOS transgenic, dystrophin-deficient mice (nNOS Tg+/mdx mice) (Fig. 1C). NOS activity assays showed that relative NO production levels generally exhibited the same trends as NOS concentration in each line assayed, although the magnitudes of the differences were smaller. Activity data were expressed relative to nNOS Tg– hearts, for which the mean value was set at 100% (SEM=5.2; n=9). Hearts of nNOS Tg+ mice had 51% more NOS activity (SEM=16.2; n=5) than nNOS Tg– hearts and nNOS Tg+/mdx hearts had 18% more activity (SEM=2.99; n=6) (Fig. 1D). However, NOS activity in nNOS Tg– hearts did not differ significantly from activity in nNOS Tg–/mdx hearts (101%; SEM=3.7; n=5).



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  		Figure 1. Expression of an active nNOS transgene in cardiac tissue. (A) Whole heart homogenates show that endogenous nNOS expression does not vary between adult mdx and C57 mice. Left three lanes are mdx mice; right three lanes are C57 mice. (B) Whole heart homogenates from nNOS Tg+ mice show a 9.4-fold increase in nNOS expression. Left three lanes are nNOS Tg– hearts; right three lanes are nNOS Tg+ hearts. (C). nNOS transgene expression in hearts of nNOS Tg+ mice is twice the expression level in nNOS Tg+/mdx hearts. Left three lanes are nNOS Tg+/mdx hearts; right three lanes are nNOS Tg+ hearts. (D). Whole heart homogenates from nNOS Tg+ mice (n=5) show a 51% increase in total NOS activity, whereas nNOS Tg+/mdx hearts (n=6) show an 18% increase in total NOS activity when compared with nNOS Tg– control hearts (n=9). Total NOS activity did not differ between nNOS Tg– hearts and nNOS Tg–/mdx hearts (n=5). Asterisk indicates significant difference from nNOS Tg– controls at P<0.05. Error bars represent SEM.

 
Dystrophin-deficient mice experience myocarditis that is reduced with increased NO production
Histological analysis showed no evidence of inflammation of nNOS Tg+ or nNOS Tg– hearts at any age sampled in mice from 3 months to 18 months of age. However, nNOS Tg–/mdx hearts contained increased numbers of inflammatory cells when compared with nNOS Tg– hearts at all ages examined. In nNOS Tg–/mdx hearts from mice ~12 months of age, macrophages were increased by 497% (nNOS Tg–/mdx=1110 cells/mm3, SEM=154; nNOS Tg–=223 cells/mm3, SEM=41), eosinophils were increased by 342% (nNOS Tg–/mdx=222 cells/mm3, SEM=27; nNOS Tg–=65 cells/mm3, SEM=14), CD4+ T-cells were increased by 1046% (nNOS Tg–/mdx=251 cells/mm3, SEM=50; nNOS Tg–=24 cells/mm3, SEM=10) and CD8+ T-cells were increased by 438% (nNOS Tg–/mdx=464 cells/mm3, SEM=14; nNOS Tg–=106 cells/mm3, SEM=27) (Fig. 2). Macrophages, the predominant inflammatory cell type present in mdx hearts, were significantly reduced in concentration by cardiac expression of the nNOS transgene. nNOS Tg+/mdx mice exhibited an 88% reduction in macrophage concentration (128 cells/mm3, SEM=26) that lowered the concentration of macrophages to levels not statistically different from dystrophin-expressing controls. The concentrations of other inflammatory cell types analyzed were not affected by the nNOS transgene, suggesting a specific effect of NO upon macrophages.



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  		Figure 2. Mdx cardiac inflammation is decreased with nNOS transgene expression. Concentrations of inflammatory cells in heart tissue. Macs, macrophages; Eos, eosinophils; CD4+, CD4+ T-cells; CD8+, CD8+ T-cells. n=5 animals per group. Asterisk indicates significant difference from controls at P<0.05. Error bars represent SEM.

 
Macrophages lyse cardiomyocytes in vitro, although there is little membrane lysis in vivo
Macrophage-mediated lysis of dystrophin-deficient, skeletal muscle fibers plays a major role in promoting the pathology of mdx skeletal muscle (33Go). We tested whether cardiac macrophages could also contribute to lysis of mdx cardiomyocytes by performing co-culture assays using primary cardiomyocytes and freshly isolated macrophages and to test whether the concentration of macrophages present in mdx hearts is sufficient to cause cardiomyocyte lysis. We observed a dose-dependent cytotoxic-response in cardiomyocyte lysis with increasing concentrations of macrophages (Fig. 3A). At a macrophage concentration comparable to that observed in nNOS Tg–/mdx hearts (1000 cells/mm3), we observed lysis of 19.4% of cardiomyocytes, showing that cardiomyocytes are susceptible to lysis by activated macrophages.



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  		Figure 3. Macrophages in mdx myocardium are not potently cytolytic. (A) Activated macrophages can lyse cultured myocardial cells. Striped bar represents concentration of macrophages found in nNOS Tg–/mdx hearts. Results are the mean value of three experiments. Error bars represent SEM. (B). Representative micrograph of a nNOS Tg–/mdx heart section stained with anti-mouse IgG showing an area of cells with membrane damage (brackets). Bar=70 µm. (C). IgG-positive areas from heart sections expressed per total section area. Black bar for nNOS Tg– mice is too small to appear in histogram. n=5 mice per group.

 
Then, we tested whether the reduction of macrophage concentrations that resulted from nNOS transgene expression in the mdx myocardium decreased cardiomyocyte membrane lysis in vivo. Cardiomyocytes with damaged cell membranes in vivo were identified immunohistochemically by the presence of IgG in the muscle cell cytoplasm (Fig. 3B). No myocytes containing IgG were observed in dystrophin-expressing hearts. Areas occupied by IgG-containing myocytes were present in nNOS Tg–/mdx hearts, but were only 1.1% of the total tissue area (SEM=0.23) (Fig. 3C). Areas containing damaged myocytes were also detectable from nNOS Tg+/mdx mice and did not differ significantly from nNOS Tg–/mdx mice (0.7% of total area; SEM=0.18). Thus, cardiomyocyte membrane damage is a minor feature of the pathology of dystrophin-deficient hearts, which is not reduced significantly by nNOS transgene expression or by decreased macrophage concentrations. The lower levels of myocyte membrane damage observed in vivo than in cytotoxicity assays suggest that mdx cardiac macrophages are less cytolytic than peritoneal macrophages that are activated in vitro.

Fibrosis is prevented in dystrophin-deficient hearts expressing an nNOS transgene
The total connective tissue content of hearts was quantified by measuring hydroxyproline concentration (52Go). We found elevated fibrosis in nNOS Tg–/mdx hearts which contained 60% more connective tissue than those in normal controls (nNOS Tg–/mdx=4.46 µg hydroxyproline/mg tissue, SEM=0.39; nNOS Tg–=2.78 µg hydroxyproline/mg tissue, SEM=0.17) (Fig. 4). Hearts from nNOS Tg+/mdx mice were significantly less fibrotic than nNOS Tg–/mdx hearts (2.86 µg hydroxyproline/mg tissue, SEM=0.33) and did not differ significantly from connective tissue concentration in control, nNOS Tg– hearts. Similarly, hydroxyproline concentration in nNOS Tg+ hearts (2.46 µg/mg; SEM=0.27) did not differ significantly from the levels in nNOS Tg– or nNOS Tg+/mdx hearts.



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  		Figure 4. Cardiac fibrosis is prevented in mdx mice with a nNOS transgene. Concentration of connective tissue in hearts. n=5 mice per group. Asterisk indicates significant difference from controls at P<0.05. Error bars represent SEM.

 
Immunohistological analysis of collagen in nNOS Tg–/mdx heart tissues showed large, fibrotic lesions composed of collagen types I, III, IV and V, which were scattered throughout the ventricles (Fig. 5). In contrast, the distributions and concentrations of each collagen type in nNOS Tg+/mdx myocardia closely resembled nNOS Tg– myocardia, with no large fibrotic lesions. No fibrotic lesions were observed in the myocardia of nNOS Tg+ mice.



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  		Figure 5. Representative micrographs showing the distribution of connective tissue and collagen isoforms in heart tissue. CI, collagen type 1; CIII, collagen type 3; CIV collagen type 4 and CV, collagen type 5. Bar=70 µm.

 
DMD-like ECG abnormalities and cardiac autonomic dysfunction in dystrophin-deficient mice are mitigated by an nNOS transgene
We observed ECG abnormalities in nNOS Tg–/mdx mice, which are characteristic of DMD patients. ECG tracings from our nNOS Tg–/mdx mice displayed deep Q-waves (–159 µV versus –61 µV in controls), a diminished S:R ratio (32 versus 54% in controls), polyphasic R-waves and frequent arrhythmias (8.7 versus 2.6/h) (Figs 6–8 and Table 1). The overwhelming majority of the arrhythmias were premature ventricular contractions (PVCs). The nNOS transgene had impressive beneficial effects as nNOS Tg+/mdx mice exhibited normalized Q-waves (–56 µV), an improved S:R ratio (38%), complete absence of polyphasic R-waves and a frequency of arrhythmias that was reduced to control levels (2.7/h). All intervals analyzed (P–R, Q–R–S, Q–T and R–R) were similar between nNOS Tg–, nNOS Tg–/mdx and nNOS Tg+/mdx mice. Analyses of the same parameters in nNOS Tg+ mice yielded values comparable to nNOS Tg– mice, although the frequency of the PVC type of arrhythmias far exceeded nNOS Tg– levels. Although there was substantial variability in PVC frequency between individual nNOS Tg+ mice, the transgene clearly had an adverse effect, suggesting that over-production of NO could lead to misregulation of cardiac function.



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  		Figure 6. ECG tracings from mdx mice show DMD-like abnormalities that are ameliorated with cardiac expression of a nNOS transgene. Representative ECG tracings collected using implantable telemetry devices. (A) Normal tracing from a nNOS Tg– mouse. (B) nNOS Tg–/mdx tracing shows deep Q-waves, peaked R-waves and diminished S-waves. (C) nNOS Tg–/mdx tracing showing polyphasic R-waves. (D) Tracing from nNOS Tg+/mdx mouse showing normalized Q-waves, R-waves and S-waves. Bar=100 ms.

 


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  		Figure 8. Mdx mice experience frequent PVCs. Representative ECG tracing from a nNOS Tg–/mdx mouse showing PVCs (arrows) which are also commonly observed in DMD patients. Bar=100 ms.

 

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  		Table 1. ECG parameters in NOS Tg–, NOS Tg–/mdx, NOS Tg+/mdx and NOS Tg+ mice
 
Heart rate variability (HRV) is an index of cardiac autonomic function, with depressed HRV indicating cardiac autonomic dysfunction (53Go). Time domain analysis showed that nNOS Tg–/mdx mice exhibited decreased HRV (3.7 versus 6.4% in controls, coefficient of variance, CV) demonstrating autonomic abnormalities (Fig. 9). HRV was more normal in nNOS Tg+/mdx mice (5.3% CV) suggesting that cardiac autonomic function was somewhat normalized with the nNOS transgene and that the dysfunction observed in nNOS Tg–/mdx hearts is related to a deficiency of NO. HRV in nNOS Tg+ mice was not statistically different from nNOS Tg– mice controls.



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  		Figure 9. HRV, an indicator of cardiac autonomic function, is normalized in nNOS Tg+/mdx mice. n=3 mice per group. Asterisk indicates significant difference from controls at P<0.05. Error bars represent SEM.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Our data show that increasing nNOS expression in dystrophin-deficient hearts has significant beneficial effects. The most striking and promising finding was that nNOS transgene expression prevented fibrosis in dystrophin-deficient hearts. Not only did the hearts of nNOS Tg+/mdx mice appear more histologically normal with an absence of large, fibrotic lesions that are prevalent in mdx hearts, but also these histological improvements were associated with improvements in physiological indices that were measured by ECG. The ECG abnormalities attributed to fibrosis of the myocardium (deep Q-waves and a reduced S:R ratio) (8Go,10Go) were completely or partially normalized. Furthermore, the polyphasic R-waves that are commonly seen in mdx mice (23Go) (present study) were absent in nNOS Tg+/mdx mice. Polyphasic R-waves likely indicate a conduction abnormality such as conduction slowing or right bundle branch block (10Go). Normalization of this feature in the nNOS Tg+/mdx mice suggests that the conduction pathway was protected from fibrosis by a mechanism similar to that protecting the myocardium, or perhaps that NO produced in transgenic hearts normalized conduction via a different mechanism. In agreement with previously published ECG data from 12-month-old mdx mice (23Go), we did not observe any other conduction defects as measured by interval durations. Our findings showing that NO-mediated prevention of fibrosis in mdx hearts results in clinically detectable improvements in cardiac function emphasizes the important role of fibrosis in the disease and suggests the possibility for new therapeutic strategies.

The reduction in myocardial fibrosis in nNOS Tg+/mdx hearts corresponds with the reduction in myocardial macrophage concentrations, although we cannot definitively conclude that mdx myocardial fibrosis is a consequence of inflammation. However, that interpretation of the data is consistent with the results of numerous investigations, which have demonstrated that NO can inhibit inflammation and that inflammatory cells can promote tissue fibrosis. NO can reduce the expression of adhesion molecules, which mediate leukocyte interactions with the vascular endothelia that precede extravasation (54Go–56Go). Conversely, NOS inhibition increases adhesion of leukocytes to vascular endothelial cells and can thereby promote extravasation of inflammatory cells (57Go). After extravasation, inflammatory cells can stimulate connective tissue synthesis through any of several, cytokine-mediated pathways that elevate the expression of collagen and other extracellular matrix molecules as part of a normal tissue repair response (38Go). Many of those profibrotic cytokines, which can be synthesized by inflammatory cells, are expressed at high levels in dystrophin-deficient muscle. In particular, TGF-ß has been implicated in promoting fibrosis in mdx and DMD muscles (40Go,42Go–44Go).

The observation that nNOS transgene expression caused large reductions in macrophage concentrations in dystrophin-deficient hearts without reducing the concentrations of CD8+ T-cells or CD4+ T-cells suggest that macrophages play a central role in any fibrosis, which is induced by inflammatory cells in mdx hearts. However, previous investigators have shown that mdx mice crossed onto the nu/nu background so that they lacked T-cells, also showed great reductions in mdx cardiac fibrosis (28Go). Together, these findings indicate that macrophages and T-cells may participate in the same pathological, fibrotic pathway in the mdx myocardium. This possibility is supported by reports showing that macrophages isolated from nu/nu mice can be less responsive to activation. For example, lipopolysaccharide stimulation of nu/nu peritoneal macrophages induced less prostaglandin E2 (PGE2) production than occurred in wild-type macrophages (58Go). Because PGE2 can stimulate mast cell degranulation (59Go) and mast cells have been implicated in the pathology of dystrophin-deficiency (60Go,61Go), lower levels of PGE2 production by macrophages in mdx-nu/nu mice may affect dystrophinopathy. On the other hand, macrophages from nu/nu mice have also been reported to reside in tissues in a more highly activated state than occurs for macrophages in wild-type mice (62Go–64Go). Thus, absence of T-cells in nu/nu mice affects basal levels of macrophage activity and their response to further activation, although whether loss of T-cell regulation of macrophage function in mdx-nu/nu mice increases or decreases fibrosis is unknown.

Manipulation of NO production in dystrophin-deficient hearts can also influence cardiomyopathy through processes, which are not related to fibrosis. NO is an important regulator of cardiac autonomic function through direct, agonistic effects upon the parasympathetic pathway and through indirect, inhibitory effects upon the sympathetic pathway, which collectively serves to protect against ventricular arrhythmias (65Go). The deleterious effects of disrupted NO production in the heart are evident in animal models where NOS inhibition decreased parasympathetic function (66Go–68Go), decreased HRV (69Go) and increased ventricular arrhythmias (70Go). Furthermore, modest overexpression of eNOS in cardiac tissue increased vagal drive and appeared to be anti-arrhythmogenic (71Go). Similarly, DMD patients experience decreased parasympathetic drive, increased sympathetic drive, decreased HRV, sinus tachycardia and an increased frequency of ventricular arrhythmias (most prominently, PVCs), all of which may reflect disruptions in cardiac NO production (14Go,15Go,17Go,19Go,20Go,72Go,73Go). We observed that features consistent with DMD-associated cardiac autonomic dysfunction such as decreased HRV and an increased frequency of PVCs in dystrophin-deficient mice and found that these abnormalities were palliated in mdx mice expressing the nNOS transgene. These data further support the hypothesis that the cardiac autonomic dysfunction associated with dystrophin-deficiency is attributable, at least in part, to disruptions in NO production. However, autonomic dysfunctions that occur in hearts of nNOS–/–mice (26Go) are not identical to those that occur in mdx hearts, which indicates that the impaired autonomic function in mdx hearts is not simply attributable to a reduction of nNOS-derived NO.

The beneficial effects of increased nNOS activity in mdx mice that express the nNOS transgene may be highly dependent on the level of nNOS-derived NO production. Expression of the transgene in mdx hearts in which endogenous levels of nNOS activity have been shown to be reduced by ~80% (23Go), yielded levels of total NO production that were 18% higher than levels measured in healthy, nNOS Tg– hearts. Our findings show that this 18% increase was associated with normalization of cardiac function. However, we also observed that nNOS transgene expression in dystrophin-expressing myocardia, in which endogenous nNOS-derived NO and transgenic nNOS-derived NO were both produced, yielded an increase in total NO production of 51%. This greater elevation of NO production was associated with a tremendous increase in the rate of PVCs, so that they occurred hundreds of times more frequently than in nNOS Tg– hearts. We observed no increase in myocardial inflammation or fibrosis in nNOS Tg+ hearts, suggesting that the elevation of PVCs resulted from disruption of autonomic function.

Sinus tachycardia, which may be caused by impaired parasympathetic function, is frequently observed in DMD patients (9Go,10Go,13Go,14Go,72Go,73Go) and is potentially influenced by NO levels in the heart. However, most reports show that NO has a positive chronotropic effect on mammalian hearts (70Go,74Go,75Go), which suggests that reductions in nNOS activity do not cause tachycardia in DMD hearts. The positive chronotropic effect of NO on heart rate may reflect a direct stimulatory effect of NO on cardiomyocytes, independent of the parasympathetic system, since the application of NO donors to excised atria in vitro can increase spontaneous beating (76Go). We did not observe an increased heart rate in mdx mice, although other investigators reported increased (23Go,26Go) or decreased (24Go) heart rates in mdx mice. However, heart rate data are subject to differences between experimental treatments of mdx mice since autonomic function is affected by the use of anesthetics (77Go) and mdx heart rate response is thought to be perturbed under stressful conditions (24Go). The use of non-anesthetized animals that were housed long-term in the cages where data were collected telemetrically provides data that are most representative of the normal heart rates of the mice.

Although disruption of nNOS function contributes to the pathology of dystrophin-deficient hearts and skeletal muscle, the relationships between dystrophin-deficiency and nNOS defects differ distinctly in skeletal muscles and hearts. In skeletal muscle, loss of dystrophin causes a large reduction in the concentration of nNOS protein and mRNA, which results in a decrease by >70% in nNOS activity in mdx muscle (46Go). However, results of the present study show that nNOS concentration is unaffected by dystrophin-deficiency in the myocardium, although nNOS activity in the dystrophin-deficient heart is reduced by ~80% (23Go). These observations indicate that dystrophin-deficiency causes defects in nNOS regulation rather than defects in nNOS expression in the heart. Perhaps, these distinct relationships between dystrophin-deficiency and nNOS dysfunction in skeletal and cardiac muscles reflect a lack of interaction between nNOS and the dystrophin complex in cardiac muscle. Unlike skeletal muscle, where nNOS and the dystrophin complex are co-distributed at the surface membrane and from which they co-fractionate (45Go), nNOS and dystrophin distributions in cardiomyocytes are not identical. Immunohistochemical observations show that dystrophin is enriched in T-tubules and at the sarcolemma of cardiomyocytes (78Go), but nNOS has been localized to the sarcoplasmic reticulum and mitochondria in addition to the sarcolemma in cardiomyocytes (47Go–50Go). Furthermore, we are not aware of any published findings showing that nNOS associates with the dystrophin complex in the myocardium.

The results of the present investigation indicate that the protective effect of nNOS transgene expression in the mdx heart may occur through its anti-inflammatory function on macrophages, as previously observed in skeletal muscle (33Go). However, our findings also show that the role of macrophages in promoting pathology differs between mdx skeletal and cardiac muscles. For example, the lysis of mdx skeletal muscle fibers is a major feature of dystrophinopathy that is largely attributable to cytolytic functions of macrophages and is greatly reduced by elevated NO production (33Go). However, there is little evidence of myocyte membrane lysis in mdx hearts, even in regions of inflammation, and elevated NO production has no significant effect on the levels of myocyte membrane damage (present study). In addition, reductions of NO production in dystrophin-deficient skeletal muscle can promote muscle pathology by increasing muscle ischemia (79Go,80Go), but mdx mice have normal cardiac vascular perfusion (35Go,81Go) and vascular irregularities are rarely reported in hearts of DMD patients (9Go,10Go,12Go,15Go).

Collectively, these findings show that perturbations in myocardial nNOS activity can contribute to the pathology of dystrophin-deficiency and suggest that manipulation of cardiac NO production may provide a therapeutic strategy for reducing cardiomyopathy in dystrophin-deficiency. However, much needs to be learned concerning the subtle but important mechanisms through which cardiac nNOS function and distribution are regulated, and merely increasing NO production by the dystrophic myocardium may increase cardiomyopathy if production levels substantially exceed normal physiological levels of production.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Animals
All animal experimentation was conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and the University of California, Los Angeles Institutional Animal Care and Use Committee. Mice were obtained from our breeding colonies housed at the UCLA vivarium with original C57BL/6 and mdx breeding pairs purchased from The Jackson Laboratory (Bar Harbor, ME, USA). Transgenic mice were generated as described previously (33Go) with the line used in the current study showing nNOS transgene expression in cardiac tissue (82Go). Mdx mice were crossed with nNOS Tg+ mice to produce dystrophin-deficient, nNOS Tg mice (nNOS Tg+/mdx mice) according to the breeding strategy described previously (33Go). Null mutation for dystrophin was confirmed by ARMS–PCR (83Go) and over-expression of nNOS was confirmed by western blot analysis. Unless otherwise noted, mice used were of mixed gender and were 12–14 months of age to ensure the presence of cardiomyopathic features in dystrophin-deficient lines.

Western blot analysis of cardiac nNOS expression
Following isoflurane-induced euthanasia, hearts were rapidly excised, rinsed in phosphate-buffered saline (PBS), pH 7.4 (140 mM NaCl, 2.7 mM KCl, 5.4 mM Na2HPO4•7H2O and 1.8 mM KH2PO4) and frozen in liquid nitrogen. Frozen hearts were homogenized in 40 volumes sodium dodecylsulfate–polyacrylamide gel electrophoresis (SDS–PAGE) reducing buffer (80 mM Tris–HCl pH 6.8, 0.1 M dithiothreitol (DTT), 70 mM SDS and 1.0 mM glycerol) with protease inhibitor cocktail (Sigma, St Louis, MO, USA). Samples were steamed for 1 min and centrifuged for 1 min at 12 000g and 4°C. Total protein concentration of the supernatant was determined by measuring absorbance at 280 nm and 80 µg of each sample was separated on 10% SDS–PAGE gels according to Laemmli (84Go). Proteins were transferred elecrophoretically onto nitrocellulose membrane, while immersed in transfer buffer (39 mM glycine, 48 mM Tris, 0.037% SDS, 20% methanol) (85Go). Uniformity of loading and transfer efficiency was confirmed by staining with 0.1% Ponceau S (Sigma). Non-specific binding was blocked by incubating membranes in blocking buffer (0.5% Tween-20, 0.2% gelatin and 3.0% dry milk) for at least 1 h. Protein samples were probed with polyclonal rabbit anti-nNOS antibody (generous gift from Dr B.S. Masters, University of Texas Health Science Center, TX, USA) for 2 h followed by horseradish peroxidase (HRP)-conjugated anti-rabbit IgG (Amersham, Piscataway, NJ, USA) for 1 h. Between incubations a series of washes in 0.1% Tween in PBS were performed. Bands were visualized via enhanced chemiluminescence (Amersham).

NOS activity assay
Total NOS activity in hearts was determined by measuring the conversion of [3H]-L-arginine to [3H]-L-citrulline using a modification of the procedure described by Bia et al. (23Go). Frozen hearts were ground under liquid nitrogen with a mortar and pestle and the resulting powder was homogenized in five volumes of 50 mM Tris pH 7.5 containing 1 mM ethylenediamine tetraacetic acid (EDTA), 1 mM ethylene glycol-bis(ß-amino ether)-N,N,N',N'-tetraacetic acid (EGTA), 1 mM DTT and protease inhibitor cocktail (Sigma). After centrifuging the homogenate at 1500g and 4°C for 10 min, 60 µl of the supernatant was incubated in 200 µl of reaction buffer with a final concentration of 50 mM Tris pH 7.5, 5 mM CaCl2, 1 mM MgCl2, 14 µM tetrahydrobiopterin, 10 µg/ml calmodulin, 4 µM flavin adenine dinucleotide, 4 µM flavin adenine mononucleotide, 1 mM reduced nicotinamide adenine dinucleotide phosphate (NADPH) and 1 µl of 1 mCi/ml [3H]-L-arginine. After a 30 min incubation at 37°C, the reactions were stopped with 20 mM sodium acetate pH 5.5, 0.2 mM EGTA, 1 mM L-citrulline and 2 mM EDTA and poured over Dowex-50W columns (Biorad, Hercules, CA, USA) previously converted to Na+ form to separate the [3H]-L-arginine from the [3H]-L-citrulline. The eluate was mixed with scintillation cocktail (Packard Bioscience, Meriden, CT, USA) and the samples counted in a LS-3133T scintillation counter (Beckman, Fullerton, CA, USA). Scintillation counts were normalized to total protein content of the homogenate as determined by measuring the absorbance at 280 nm and expressed as a percent of control values.

Histology
Following euthanasia with isoflurane, hearts were rapidly excised, transferred to liquid nitrogen-cooled isopentane-filled vials and stored at –80°C until use. Frozen hearts were sectioned through the short axis of the ventricles at a thickness of 10 µm. For immunostaining, sections were air-dried and fixed in acetone, and endogenous peroxidase activity was quenched with 0.03% hydrogen peroxide. Sections were incubated in PBS with 2% gelatin and 3% bovine serum albumin to prevent non-specific binding of antibodies. Tissues were incubated with primary antibodies overnight at 4°C, host-appropriate biotinylated secondary antibodies (Vector Laboratories, Burlingame, CA, USA) for 30 min at room temperature, and HRP-avidin D for 30 min. Sections were washed with PBS between antibody incubations. Staining was visualized using 3-amino-9-ethyl carbazole (AEC, red) (Vector) as substrate. Antibodies used for inflammatory cell staining were rat monoclonal anti-mouse F4/80 (for macrophages), anti-mouse CD4 and anti-mouse CD8, all obtained from supernatants of hybridoma cultures (hybridomas obtained from American Type Culture Collection, Bethesda, MD, USA) and polyclonal rabbit anti-murine eosinophil granule major basic protein (a generous gift from Dr J.J Lee, Mayo Clinic Scottsdale, AZ, USA) (86Go). The concentrations of inflammatory cells were measured by histomorphometry as previously described (87Go).

Collagen staining was performed as described earlier with a few exceptions. Sections were incubated with primary antibodies for 3 h at room temperature and with host-appropriate fluorescent-conjugated secondary antibodies (Vector) for 1 h. Primary antibodies used were rabbit anti-rat collagen type I (Chemicon, Temecula, CA, USA), rabbit anti-rat collagen type III (Chemicon), goat anti-human collagen type IV (Southern Biotech, Birmingham, AL, USA) and goat anti-human collage type V (Southern Biotech). Sections stained for collagen were qualitatively analyzed for the presence of foci of fibrosis.

IgG staining
Cardiomyocytes with damaged membranes were identified by IgG staining as performed previously by Hainsey et al. (27Go). Transverse sections through the ventricles, 10 µm in thickness, were incubated for 30 min with a 1% gelatin solution in PBS to block non-specific binding. After rinsing with PBS, the sections were incubated with FITC-conjugated mouse anti-IgG (Vector Laboratories) for 1 h. A series of three PBS washes were performed and the sections were viewed and analyzed on a microscope equipped with fluorescent optics. Areas of damaged cells were quantified according to the previous technique (88Go); a sampling grid was superimposed on the tissue and the number of intercepts that overlie cell IgG-positive cells were counted and expressed as a percentage of the total number of intercepts.

Cytotoxicity assays
Primary cardiac cultures were generated from neonatal C57 mice according to the modified protocol of Sen et al. (89Go). Hearts were removed from 2-day-old C57 mice and trimmed of connective tissue and atria. Cardiomyocytes were dispersed from the ventricles by digestion with 0.2% collagenase II (Invitrogen, Carlsbad, CA, USA) and 0.6% pancreatin (Sigma) in a buffer containing 0.8 mM MgSO4, 116 mM NaCl, 0.5 mM KCl, 11 mM NaH2PO4, 5.5 mM glucose and 20 mM HEPES. The cell suspension was preplated twice for 30 min each to enrich for cardiomyocytes. Cells were counted and the enriched cardiomyocytes were plated in 4:1 DMEM/medium 199 (Sigma) containing penicillin and streptomyocin and supplemented with 10% FBS (Omega Scientific, Tarzana, CA, USA) at 1800 cells/mm2 which produced a confluent layer in gelatin-coated 96-well plates. The cultures were maintained at 37°C with 5% CO2 and began beating synchronously 2–3 days following plating.

Macrophages for co-culturing were obtained by modification of our previously described technique (90Go). Peritoneal exudate was collected from adult C57 mice 3 days following intraperitoneal injection of 12% sodium caseinate. The exudate was filtered through 70 µm nylon mesh, cells pelleted at 500g and resuspended in 0.85% NH4Cl to lyse erythrocytes. The cells were re-pelleted, resuspended in HBSS and overlaid onto Histopaque 1077 (Sigma). After centrifuging at 500g for 45 min the purified macrophages were collected from the interface and purity of the population was determined histologically to be >90%.

Approximately 5 days after plating the cardiomyocyte cultures, the cells were loaded with 51Cr (as previously described) (33Go) and co-cultured for 16 h with varying concentrations of freshly isolated macrophages activated with 0.6 µM PMA. The medium was then assayed for 51Cr release by scintillation counting. Cytotoxicity was expressed as a percentage of total lysis by setting 0% as 51Cr released spontaneously by cardiomyocytes incubated without macrophages. The 51Cr release into the medium by cardiomyocytes lysed with 0.1% Triton X-100 (Sigma) was determined to establish 100% cytotoxicity.

Hydroxyproline assay
Total connective tissue content of hearts was quantified by measuring the amount of hydroxyproline concentration in the tissues according to the technique of Kivirikko et al. (52Go) that we have used previously (91Go).

Electrocardiography
ECG data was collected in awake, freely moving mice by implanting radio telemetry devices (TA10ETA-F20, Transoma-Data Science Intl. (DSI), St Paul, MN, USA). Measuring ECG activity in conscious mice is advantageous since the confounding effects of anesthesia upon cardiac function are obviated. Transmitter units were implanted in the peritoneal cavity of anesthetized mice and the two electrical leads were secured near the apex of the heart and the right acromion in a lead II orientation. Mice were housed singly in cages over antenna receivers connected to a computer system for data recording. Unfiltered ECG data was collected for 10 s each hour for 35 days. The first 7 days of data were discarded to allow for recovery from the surgical procedure and ensure any effects of anesthesia had subsided. Data waveforms and parameters were analyzed with the DSI analysis packages (ART 3.01 and Physiostat 4.01) and measurements were compiled and averaged to determine heart rates, ECG wave heights and interval durations. Raw ECG waveforms were scanned for arrhythmias by two independent observers.

Time-domain measures of HRV
The CV (%), an index of HRV, was determined from sequential R–R intervals as previously described by Gehrmann et al. (53Go). Briefly, the standard deviation of all normal R–R intervals was divided by mean of all R–R intervals: SDNN/RRmeanx100.

Statistics
Statistical significance of differences between groups was determined using the two-tailed Student's t-test. The P-value was set at 0.05. ANOVA was used for four-way analysis of cardiac telemetry data.


   ACKNOWLEDGEMENTS
 
The authors thank Dr Joshua Goldhaber for helpful discussions on the ECG data analysis and Helen C. Chang for assistance in ECG data analysis. This work was supported by grants from the American Heart Association 0325146Y (M.W.H.), the National Institutes of Health AR40343 (J.G.T.), AR47721 (J.G.T.), the Muscular Dystrophy Association (J.G.T.) and the Laubisch Endowment (K.P.R.).

Conflict of Interest statement. None declared.



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  		Figure 7. Q-wave amplitude is normalized in mdx mice with cardiac expression of a nNOS transgene. n=3 mice per group. Asterisk indicates significant difference from controls at P<0.05. Error bars represent SEM.

 

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 MATERIALS AND METHODS
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