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Human Molecular Genetics, 2004, Vol. 13, No. 15 1669-1675
DOI: 10.1093/hmg/ddh174
Human Molecular Genetics, Vol. 13, No. 15 © Oxford University Press 2004; all rights reserved

Full-length dystrophin expression in half of the heart cells ameliorates ß-isoproterenol-induced cardiomyopathy in mdx mice

Yongping Yue1, Jeffrey W. Skimming2, Mingju Liu1, Tammy Strawn2 and Dongsheng Duan1,*

1Department of Molecular Microbiology and Immunology and 2Department of Child Health and Department of Medical Pharmacology and Physiology, The University of Missouri, School of Medicine, Columbia, MO 65212, USA

Received April 12, 2004; Accepted May 26, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Gene therapy holds great promise for curing Duchenne muscular dystrophy (DMD), the most common fatal inherited childhood muscle disease. Success of DMD gene therapy depends upon functional improvement in both skeletal and cardiac muscle. Numerous gene transfer studies have been performed to correct skeletal muscle pathology, yet little is known about cardiomyopathy gene therapy. Since complete transduction of the entire heart is an impractical goal, it becomes critical to determine the minimal level of correction needed for successful DMD cardiomyopathy gene therapy. To address this question, we generated heterozygous mice that persistently expressed the full-length dystrophin gene in 50% of the cardiomyocytes of mdx mice, a model for DMD. We questioned whether dystrophin expression in half of the heart cells was sufficient to prevent stress-induced cardiomyopathy. Heart function of mdx mouse is normal in the absence of external stress. To determine the therapeutic effect, we challenged 3-month-old mice with ß-isoproterenol. Cardiomyocyte sarcolemma integrity was significantly impaired in mdx but not in heterozygous and C57Bl/10 mice. Importantly, in vivo closed-chest hemodynamic assays revealed normal left ventricular function in ß-isoproterenol-stimulated heterozygous mice. Since the expression profile in the heterozygous mice mimicked viral transduction, we conclude that gene therapy correction in 50% of the heart cells may be sufficient to treat cardiomyopathy in mdx mice. This finding may also apply to the gene therapy of other inherited cardiomyopathies.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Duchenne and Becker muscular dystrophy (DMD, BMD) are among the most common single gene disorders, affecting approximately one in every 3500 newborn boys and one in every 20 000 newborns, respectively (1). Genetic defects in the dystrophin gene are responsible for DMD and BMD. Mutations that completely eliminate the dystrophin protein expression result in lethal childhood onset DMD. Abnormalities in the level and/or structure of the dystrophin protein are often associated with much milder allelic BMD. Despite the fact that the dystrophin gene is the first disease-related gene identified by positional cloning in 1987 (2), limited progress has been made to improve the clinical outcome of DMD and BMD patients. Cardiac dysfunction plays a major role in both morbidity and mortality of patients with DMD and BMD. More than one-tenth of the DMD patients die of heart failure (3). More than 90% of BMD patients have cardiac phenotypes (4). Furthermore, in a subset of patients with dystrophin gene mutations, the only clinical manifestation is the allelic X-linked dilated cardiomyopathy (5). Other than heart transplantation, symptom-relieving medicine (such as angiotensin-converting enzymes and ß-blockers) is currently the only available treatment (6).

Replacing and/or repairing the mutated gene by gene therapy represents a promising cure. In the last decade, numerous advances have been made in DMD skeletal muscle disease gene therapy (7). For example, to circumvent the size limitation of the 2.4 Mb dystrophin gene, novel C-terminal truncated microgenes were developed. These microgenes have been shown to improve skeletal muscle morphology and contractility (8,9). Furthermore, it has been demonstrated that as little as 20% of the normal dystrophin concentration is sufficient to avoid myopathy in limb muscle and the diaphragm (10,11).

Unlike multinucleated skeletal muscle cells, intercalated discs separate cardiomyocytes. As a result, viral transduction is restricted to each individual heart cell, rather than diffusing through entire fiber (12–14). This structural difference necessitates establishment of heart specific parameters for cardiac gene therapy. We recently reported that micro-dystrophin could restore the dystrophin–glycoprotein complex and maintain sarcolemma integrity in the heart of mdx mice, a mouse model for DMD (14). Yet, the number of dystrophin expressing cells needed for cardiomyopathy therapy has not been determined.

In this study, we used 3-month-old female heterozygous mice, F1 from C57Bl/10 (BL10) and mdx crossing, as an experimental model to evaluate whether persistent full-length dystrophin expression in half of the cardiomyocytes was sufficient to improve heart function in mdx mice. Generally mdx mice exhibit a normal cardiac function. However, the myocardium of mdx mice is extremely vulnerable to mechanical stress (15). To determine whether non-uniform dystrophin expression in ~50% of the cardiomyocytes was sufficient to ameliorate the mdx cardiomyopathy, we challenged mdx, heterozygous and BL10 mice with ß-isoproterenol. Our results suggest that partial expression of dystrophin can protect the mdx hearts from ß-isoproterenol-induced damage. Notably, cardiomyocyte sarcolemma integrity was significantly strengthened and left ventricular systolic and diastolic function was completely recovered to the normal BL10 mouse level.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Heterozygous female mdx mice as a model to evaluate therapeutic efficacy of partial dystrophin expression
As a result of random X-chromosome inactivation (16), ~50% of the heart cells are expected to be dystrophin positive. To accurately determine dystrophin expression, we manually counted all dystrophin-positive cells in each heart section (~50 000 cells for one BL10 section) and quantified at least three representative sections (base, body and apex) for each heart. As shown in Figure 1B, there were 2545±224 cardiomyocytes/mm2 in normal BL10 mice (N=3). In maternal (N=7) and paternal (N=6) heterozygous mice, there were 1304±76 and 1410±110 dystrophin-positive cells/mm2, respectively. In both the cases, about half of the heart cells stained positive for dystrophin (51.22% in maternal and 55.40% in paternal). This morphometric quantification result was further confirmed by immunoblot with whole heart lysates (Fig. 1C). In skeletal muscle, the syncytial nature of the myofiber structure allows dystrophin transcripts produced from one nucleus to diffuse through the entire fiber. Eventually, a homogenous dystrophin expression is achieved in all myofibers in skeletal muscle (Fig. 1D, top left panel). Interestingly, dystrophin-positive cells were distributed in patches in the hearts of heterozygous mice. In some areas, more than 90% cells were positive. In others, only scattered dystrophin expressing cells were seen (Fig. 1D, bottom panels). This non-uniform pattern resembles transduction profiles seen in viral gene transfer studies (12–14).



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  		Figure 1. Breeding scheme and dystrophin expression profile. (A) mdx mice were crossed with BL10 mice to generate paternal and maternal female heterozygous mice. In maternal heterozygous mice, mutated dystrophin gene is inherited from the mdx mother. In paternal, the mutated gene is from the mdx father. (B) Morphometric quantification of dystrophin-positive cardiomyocytes in the hearts. There is no significant difference between maternal and paternal mice. However, values in BL10 mice are significantly higher than those in heterozygous mice. N=3 for BL10, N=7 for maternal heterozygous, N=6 for paternal heterozygous. (C) Representative western blot of full-length dystrophin expression (427 kDa, arrow) in the hearts of different mouse strains. Hetero, heart lysate from heterozygous mouse. (D) Top panels are representative immunofluorescence staining of dystrophin expression in anterior tibialis muscle (TA, top left) and the heart (top right) of heterozygous mice. Higher power photomicrographs of two representative areas in the heart are displayed in bottom panels. Box 1 depicts a region of high dystrophin expression. Box 2 is a low dystrophin expression area. Scale bar in the top left panel (TA) is 300 µm. Scale bar in the bottom right panel (Box 2) is 100 µm. This scale bar also applies to the photomicrograph in the bottom left panel (Box 1).

 
50% Dystrophin expression reduces heart hypertrophy in mdx mice
To determine whether there are anatomic changes in the mdx heart, we examined a relatively large population of age- and sex-matched mice (32 BL10 and 40 mdx). As summarized in (Table 1), both the whole heart and the isolated ventricle were heavier in mdx mice than in BL10 mice. However, ratios of heart weight (HW) to body weight (BW) (HW/BW) and ventricular weight (VW) to BW (VW/BW) were similar between the two strains, indicating that the change in heart size was appropriate for the change in body size. Nevertheless, a dramatic change in skeletal muscle pathology caused a significant reduction in the ratios of HW to tibialis muscle weight (TW) (HW/TW) and VW to TW (VW/TW) in mdx mice. In heterozygous mice, all the above indices were similar to BL10. Obviously, the lack of skeletal muscle pathology in heterozygous mice has contributed to this observation (17). Taken together, the similarity between heterozygous and BL10 HW suggests that the threshold for normal cardiac anatomy has been met with half of the heart cells expressing dystrophin.


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  		Table 1. Experimental mice characterization
 
Heterozygous mouse hearts are more resistant to stress-induced sarcolemma damage than mdx hearts
Dystrophin plays a critical role in maintaining muscle cell integrity during contraction. Muscle integrity can be measured using an in vivo Evans blue dye (EBD) uptake assay in which the dye diffuses only into injured muscle cells. Since the mdx heart functions well for normal daily activity (14,15), we pharmacologically stressed the hearts with ß-isoproterenol. As shown in Figure 2A, very few EBD-positive cells were detected in the BL10 hearts after drug administration. In sharp contrast, mechanical challenge from ß-isoproterenol resulted in 11.26±3.40% EBD-positive area in the mdx hearts. Importantly, the EBD-positive area was reduced to 2.37±0.70% in heterozygous mice. Additional immunostaining confirmed that these EBD-positive cells were not expressing dystrophin (Fig. 2B).



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  		Figure 2. Quantitative evaluation of sarcolemma integrity in the hearts. (A) Representative photomicrographs of the entire heart sections from BL10, mdx and heterozygous mice. Scale bar, 1 mm. EBD uptake is visualized under Texas Red channel. Bar graph shows the percentage of injured area (EBD-positive region) in the hearts. The results are significantly different among all groups and between any paired strains. N=17 for BL10, N=11 for mdx, N=10 for heterozygous. Hetero, heterozygous mice. (B) Representative higher magnification photomicrographs of the heart sections. Dystrophin expression was detected by indirect immunofluorescence staining (FITC channel). EBD uptake in the same field is shown in Texas Red channel photomicrographs. EBD uptake can also be visualized under FITC channel as cells that were filled up with fluorescence signal. However, this is different from continuous, peripheral membrane staining of dystrophin. Arrow, an EBD positive-region in a heterozygous mouse heart section. Scale bar, 300 µm.

 
Full-length dystrophin expression in half of the cardiomyocytes corrects hemodynamic dysfunction in the mdx hearts
The ultimate goal of DMD cardiomyopathy gene therapy is to improve heart hemodynamic function. However, mdx heart function seems to be adequate in the absence of exogenous stress (Fig. 3B). To determine whether dystrophin expression in ~50% of the heart cells could improve hemodynamic performance, we challenged the mice with ß-isoproterenol.




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  		Figure 3. In vivo hemodynamic evaluation of the heart function. (A) Representative tracing following ß-isoproterenol challenge in BL10, mdx and heterozygous mice. dP/dt, rate of pressure change in left ventricle; LVP, left ventricular pressure. (B) Hemodynamic profile in different mouse strains in the absence (open bar) or presence (filled bar) of ß-isoproterenol challenge. Developed pressure was calculated from the difference between minimal diastolic pressure and systolic pressure. For ß-isoproterenol treated group, N=10 for BL10, N=9 for mdx and N=11 for heterozygous. For non-treated group, N=3 for BL10, N=10 for mdx and N=3 for heterozygous. There is no statistical difference among BL10, mdx and heterozygous mice in the absence of ß-isoproterenol stimulation. *, ß-isoproterenol treated mdx was significantly different from similarly treated BL10 or heterozygous. #, ß-isoproterenol treated mdx was significantly different from the non-treated mdx.

 
Consistent with our EBD uptake results, mdx left ventricular function was significantly compromised following ß-isoproterenol stress (Fig. 3). Specifically, peak systolic pressure and maximal dP/dt were reduced, indicating systolic dysfunction. Impaired relaxation during diastole was reflected by an increase in the peak dP/dt minimal and minimal diastolic pressure. As a result, the left ventricular developed pressure was also substantially reduced in the mdx mice.

Although only ~50% of the heart cells were expressing dystrophin in heterozygous mice, heart function was normal in these mice even under ß-isoproterenol stress. There was no worsening of dP/dt maximal, dP/dt minimal, systolic pressure, minimal diastolic pressure and developed pressure following ß-isoproterenol challenge (Fig. 3).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Cardiomyopathy commonly precipitates death in DMD and BMD patients. Despite tremendous progress in our understanding of the pathogenesis, treatment remains enigmatic for DMD and BMD. Since the lack of dystrophin protein is primarily responsible for both skeletal muscle and the heart dysfunction (2,18), restoring dystrophin expression by gene therapy may ultimately cure this devastating disease. Successful therapy of DMD and BMD necessitates functional improvements in both skeletal and cardiac muscle. Experimentation with the murine model has revealed many important aspects of treating skeletal muscle disease in DMD. These include the determination of the minimal amount of dystrophin necessary for therapy and the development of novel dystrophin isoforms that can fit into viral vectors. In skeletal muscle, a 50-fold over-expression is not toxic (19) and as low as 20% of the normal dystrophin concentration is sufficient to maintain muscle function (10,11). Patient studies have demonstrated that an even distribution of only 50% of the normal cells was enough to avoid the most severe form of skeletal muscle injury (20,21). In a 42-year-old woman carrier, 50% mosaic expression in skeletal muscle was associated with very mild skeletal muscle weakness, yet her abnormal electrocardiogram suggested cardiomyopathy (22). Clearly, dystrophin expression in 50% of the skeletal myofibers has minimal protection on the heart. Currently, identifying the minimal range of dystrophin expression necessary for normal heart function is the top priority in DMD cardiomyopathy gene therapy.

Recent developments in gene transfer methods have allowed transduction of 50–80% of adult mouse hearts with adenovirus and AAV (12,13). To address whether dystrophin expression in only portions of the heart ameliorates stress-induced cardiomyopathy in the mdx hearts, we used the genetically-defined female heterozygous model. A mosaic dystrophin expression has been shown in female heterozygous mdx hearts (23–26). Because of random X-chromosome inactivation, ~50% of the cardiomyocytes are expected to express dystrophin in heterozygous mice (16). However, some previous studies have shown up to 70% of the dystrophin-positive cells in either maternal (24) or paternal (26) female heterozygous mice. Since <5000 heart cells were quantified per heart section in these studies, the inconsistency may have resulted from an insufficient sampling. Our comprehensive quantifications at both cellular (Fig. 1B and D) and organ levels (Fig. 1C) demonstrate that irrespective of the origin of the mutant dystrophin gene (paternal or maternal), only half of the cardiomyocytes express dystrophin in heterozygous mice.

The mdx mouse is the prototype murine DMD model. Unfortunately, its heart pathology has not been evaluated systematically. Two studies suggest that mdx hearts may be hypertrophic (15,27). However, the sample sizes (five and 12 mice, respectively) were too small in these studies to reveal a statistically significant hypertrophy. In this study, we characterized mdx cardiomyopathy. Although we found a significant increase in HW and VW, this increase may simply be accounted for by a concomitant increase in BW and limb muscle weight in adult mdx mice (28,29). The most significant cardiac pathologies detected in mdx mice were the increased EBD uptake and the reduced left ventricle function following ß-isoproterenol challenge. These pathologic findings are consistent with in vitro results obtained in Langendorff heart preparations (15). In both the cases, increased heart work led to left ventricular dysfunction in mdx mice.

The protection afforded by 50% of the dystrophin-positive cardiomyocytes in heterozygous mice was first demonstrated by a normalization of HW and VW. To more accurately determine the therapeutic efficacy of this degree of dystrophin expression, we challenged the mice with ß-isoproterenol. Heart EBD uptake was dramatically decreased in the heterozygous mouse hearts. It was reduced to one-fifth of that of the mdx heart EBD uptake. How partial correction leads to global protection in dystrophin-negative cells remains unclear. Considering the fact that even in dystrophin null mdx hearts EBD leakage was detected only in ~11% of the heart tissue, it is possible that the stress we applied may not have been high enough to damage all cardiomyocytes in the heart. Alternatively, unknown compensatory mechanism(s) of the heart may have masked cardiac dysfunction. Taken together, our result suggests that a significant improvement in sarcolemma integrity has been achieved in the heterozygous mouse hearts. Since EBD uptake in heterozygous heart was still higher than that in the BL10 heart, a higher proportion of dystrophin-positive cells might provide better protection.

The most impressive correction was seen in the in vivo hemodynamic assay. A 50% of the dystrophin-positive cardiomyocytes yielded a 100% protection to ß-isoproterenol stress in heterozygous mice. Despite meaningful results obtained in dP/dt and heart pressure measurements, heart rate was not significantly altered by ß-isoproterenol challenge (Fig. 3). An immediate increase in heart rate has been demonstrated during continuous ß-isoproterenol infusion (30–32). We can only speculate that the delay from ß-isoproterenol administration to heart rate measurement accounted for the lack of an increase in heart rate in our study.

In this study, we have demonstrated the therapeutic effect of partial dystrophin expression in mdx mice. Our results suggest that a 50% correction in the mdx heart is sufficient to enhance heart cell sarcolemma integrity and normalize hemodynamic function in young adult mice. Expressing full-length dystrophin in 50% of the heart cells holds great therapeutic promise.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Breeding, heart harvesting and quantification of dystrophin expression in the heart
All animal experiments were carried out in accordance with NIH and institutional guidelines of the University of Missouri. Parental BL10 and mdx mice were purchased from Jackson Laboratory. Colonies were subsequently established by breeding at the University of Missouri. Paternal heterozygous mice were obtained by crossing mdx male with BL10 female. Maternal heterozygous mice were obtained by crossing BL10 male with mdx female (Fig. 1A). All the experiments were performed in 3-month-old female mice.

To characterize the hearts from different strains, mice were euthanized by cervical dislocation. The hearts were removed and flushed with Phosphate-buffered saline (PBS) three times and blot dried. After measuring the whole HW, the atria were removed and the combined right and left VW were measured. To determine the relative HW and VW, we also isolated left and right anterior tibialis muscle (TA) and measured TA weights. The average of both TA weights was defined as TW.

To quantify the number of dystrophin-positive cells, freshly dissected hearts were snap frozen in liquid nitrogen-cooled isopentane and indirect immunofluorescence was performed with an anti-dystrophin C-terminal antibody (Dys-2, Novocastra Laboratories Ltd, Newcastle, UK) in 8 µm cryosections according to our published protocol (14). To obtain a comprehensive evaluation, more than three representative sections from the base, the middle portion and the apex were quantified for each heart sample. A montage of the entire heart section was assembled from digitized images for each section. All the dystrophin-positive cells in the entire heart section were manually counted using an electronic colony counter. The relative heart area was determined with Scion Image 1.62 software. The absolute heart area (mm2) was deducted from the relative heart area by converting relative length unit to metric unit using a reference KR 867 micrometer (Klarmann Rulings Inc., Litchfield, NH, USA). The number of total dystrophin-positive cells was normalized by the cross-section area of the heart.

To confirm the morphometric quantification result, western blot was performed on heart lysates from BL10, mdx and heterozygous mice. Briefly, freshly dissected hearts were pulverized in liquid nitrogen and resuspended in a lysis buffer containing 5 mM ethylene-bis-oxyethylenetrinitrilo-tetraacetic acid (EGTA), 1% sodium dodecyl sulfate (SDS) and 1x protease inhibitor (Roche Applied Science, Indianapolis, IN, USA; catalog no. 1836145). After a 5 min sonication at a power output of 6.0 (Misonix Sonicator 3000, Misonix Inc., Farmingdale, NY, USA), lysates were cleared by a 5 min spin at 4°C at 10 000 rpm on a desk-top centrifuge (Eppendorf Centrifuge 5417C). An amount of 100 µg of supernatants from each strain were electrophoresed at 4°C for 15 h on a 6% SDS–polyacrylamide gel and the transferred to Hybond nitrocellulose membrane (Amersham Bioscience, catalog no. RPN303E). Protein loading and transfer were confirmed by Ponceau-S stain. Blots were first probed with Dys-2 antibody (C-terminal specific, 1 : 100 dilution) followed by an HRP-conjugated anti-mouse secondary antibody (1 : 2000, Jackson Immunoresearch Laboratories, West Grove, PA, USA). Signals were revealed by ECL immunodetection system (Amersham Bioscience, catalog no. RPN2106).

EBD uptake in the heart
Sarcolemma integrity of the cardiomyocytes was determined according to EBD uptake after ß-isoproterenol (a positive inotropic drug) challenge. EBD (10 mg/ml in PBS, 20 µl/g BW) was injected into mouse tail vein at 24 h before heart harvesting. Three dosages of ß-isoproterenol (350 ng/g BW, intra-peritoneal injection, i.p.) were sequentially administered at 12, 18 and 21 h later. EBD uptake was visualized under the Texas Red channel by a Nikon E800 fluorescence microscope. Percentage of EBD-positive area was determined with Scion Image 1.62 software.

Closed-chest in vivo hemodynamic assay
To determine the heart hemodynamic function, we anesthetized mice with 2% isoflurane induction followed by an i.p. injection of a freshly prepared cocktail containing 720 µg/g urethane, 5 µg/g etomidate and 1 µg/g morphine (12,33). Under a dissection microscope, a midline incision was made to expose the right common carotid artery. A 1.4 F Millar micro-tip pressure catheter (Millar Instrument, Houston, TX, USA; catalog no. SPR-671) was then inserted into the right common carotid artery and advanced through the aortic valve into the left ventricle. The heart rate, systolic and diastolic pressure, and the rate of pressure change (dP/dt maximal, dP/dt minimal) were recorded at a sampling rate of 500 Hz through a BIOPAC MP100 data acquisition unit (Biopac Systems Inc., Goleta, CA, USA). An 8 s recording was analyzed with AcqKnowledge software for each mouse. Since the end of diastole was not clearly demarcated in every mouse, we opted to use minimal left ventricular pressure as an alternative index for diastolic function.

In the absence of external stress, mdx heart function was fully compensated as shown previously (15). To measure cardiac protection by dystrophin, mice were challenged with ß-isoproterenol. Briefly, three doses of ß-isoproterenol were sequentially administrated (i.p. 350 ng/g BW) at 12, 6 and 3 h before the hemodynamic assay. In these studies, the hemodynamic parameters were recorded and analyzed exactly the same way as in the absence of ß-isoproterenol challenge.

Statistical analysis
Data are expressed as mean ± SEM. We first assured that the data was normally distributed using the Shapiro–Wilk test in SAS software (SAS Institute Inc., Cary, NC, USA). We then performed one-way analysis of variance tests to determine whether the genetic differences affected each end-point. Post hoc analyses were performed using homoscedastic Student's t-tests. A significance level was set as 0.05.


   ACKNOWLEDGEMENTS
 
We thank Dr Robin L. Davisson, Dr Timothy E. Lindley, Mr James Bosanquet and Mr Vinay Rawlani for their help in developing the in vivo hemodynamic assay. We thank Dr Richard Tsika for sharing his heart harvesting technique and Ms Bin Ge for statistic analysis. We also thank Dr Markay Scott and Ms Lynne Smaile for their help. This work was supported by grants from the National Institute of Health (AR-49419, D.D.), Muscular Dystrophy Association (D.D. and J.S.) and University of Missouri Research Board (D.D.).


   FOOTNOTES
 
* To whom correspondence should be addressed at: Department of Molecular Microbiology and Immunology, The University of Missouri, School of Medicine, One Hospital Drive, Room M610G, MSB, Columbia, MO 65212, USA. Tel: +1 5738849584; Fax: +1 5738824287; Email: duand{at}missouri.edu


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

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