Duchenne muscular dystrophy is caused by defects in the dystrophin gene, and the mdx mouse is the most frequently employed genetic model of this disease. It is well known that different muscle groups do not respond in the same way to dystrophin deficiency. In particular, the mdx mouse diaphragm exhibits severe morphological and functional changes not found in other mdx muscles. Use of early generation adenoviral vectors to deliver genes to the diaphragm in immunocompetent mdx mice has been associated with substantial functional toxicity and a rapid loss of transgene expression. Here we determined the response to dystrophin gene replacement in the mdx diaphragm using a “gutted” adenoviral vector that contains the coding sequence of two full-length dystrophin genes and is deleted of most viral DNA sequences. At 1 wk postdelivery of the vector, 23.6 ± 4% of total fibers in the injected diaphragm bundle expressed dystrophin at the sarcolemma, which remained stable over the study duration of 30 days without the need for continuous immunosuppression. Treated diaphragms showed a significantly improved resistance to the abnormal force deficits induced by high-stress muscle contractions, the latter being a functional hallmark of dystrophin-deficient muscle. This functional amelioration was achieved despite the presence of mildly increased inflammation (CD4+ and CD8+ lymphocytes) within the vector-treated diaphragms. To our knowledge, this is the first demonstration that a viral vector can achieve reversal of functional abnormalities in the dystrophic diaphragm via therapeutic dystrophin gene transfer without the need for sustained immunosuppressive therapy.
- Duchenne muscular dystrophy
- gene therapy
- dystrophin deficiency
- viral vectors
- mdx mouse
duchenne muscular dystrophy (DMD) is the most common fatal X-linked recessive disorder in humans, affecting ∼1 in 3,500 live male births. The disease is caused by genetic mutations of the dystrophin gene (16), which encodes a large (427-kDa) cytoskeletal protein normally found on the cytoplasmic surface of the muscle cell membrane, or sarcolemma. The characteristic muscle pathology found in DMD patients consists of myofiber necrosis with ensuing muscle fiber loss, fibrosis, and fatty cell infiltration. This leads to progressive weakness of all skeletal muscles, including the diaphragm and other respiratory muscles. Consequently, ventilatory insufficiency is a central problem in the management of DMD patients, and most patients will die in their late teens or early twenties unless supported by mechanical ventilation (41).
Because of a lack of effective therapeutic options, DMD has been considered a prime candidate for cell or gene therapy, the goal of which is to transfer normal copies of the dystrophin gene into the muscle fibers of affected patients. To be clinically feasible, two essential features of such an intervention in patients are 1) an appropriate dystrophin gene construct, expressed at adequate levels to achieve improvements in muscle function, and 2) a lack of significant vector toxicity. To address these requirements, newer generation AdV, also referred to as helper-dependent or “gutted” vectors, have been developed. A murine model of DMD, the mdx mouse, also lacks dystrophin due to a nonsense point mutation in exon 23 of the dystrophin gene (40). We (11) and others (8, 12) have recently reported that gutted AdV are able to achieve sustained dystrophin gene expression in adult mdx mice without the need for concomitant host immunosuppression.
All studies to date that have employed gutted AdV to achieve therapeutic dystrophin gene transfer in mdx mice have limited their evaluation to the hindlimb musculature (5, 8, 11, 12, 15). However, it is important to recognize that different skeletal muscle groups do not respond in the same way to dystrophin deficiency. For example, extraocular muscles in mdx mice and humans are spared from clinical disease (38). Furthermore, hindlimb muscles of mdx mice show only mild fibrosis or functional alteration until late in life (22, 31, 43). On the other hand, the mdx mouse diaphragm exhibits major fibrosis and myofiber loss as well as greatly impaired contractile function from an early age (35, 43). Interestingly, it has also been reported that the effects of tumor necrosis factor-α gene inactivation on mdx diaphragm and quadriceps pathology are diametrically opposed (42). The precise reasons for these muscle-specific divergences in the response to dystrophin deficiency are not well understood. However, the findings are consistent with recently published data suggesting that responses of a particular muscle to a lack of dystrophin are likely determined by unique aspects of its underlying constitutive gene expression profile (37, 38).
In view of the above observations, it is evident that the response of dystrophin-deficient muscles to gene therapy may also be governed by muscle group-specific factors. Therefore, although dystrophin gene transfer with gutted AdV has been shown to be beneficial in mdx mouse hindlimb muscles, this cannot automatically be assumed to be the case for other skeletal muscle groups. In the case of the diaphragm in particular, this issue is of major importance for at least two reasons. First, there is a much greater phenotypic resemblance between human DMD muscles and the mdx diaphragm when compared with any other mdx muscle (43). Second, targeting of the respiratory muscles will ultimately be required to prolong the lifespan of DMD patients, since respiratory insufficiency is the most frequent cause of death. Accordingly, the primary objectives of the present study were 1) to evaluate the physiological efficacy of full-length dystrophin gene transfer to the mdx mouse diaphragm using a gutted AdV deleted of all viral genes, and 2) to assess the cellular immune response to AdV-mediated dystrophin gene transfer as well as its relationship to the stability of dystrophin expression in the mdx diaphragm under these conditions.
MATERIALS AND METHODS
HDCBDysM Construction and Growth
The gutted AdV used in these experiments, henceforth referred to as HDCBDysM, has been previously described in detail (11). The construct contains two full-length murine dystrophin cDNAs, both regulated by a hybrid cytomegalovirus enhancer/chicken β-actin promoter that is highly active in adult skeletal muscle (18). The vector was propagated using the 293Cre-loxP system (30) and purified by two consecutive CsCl gradient centrifugations. The CsCl was removed from the viral preparation by chromatography on Sephadex G25 columns (Amersham Pharmacia Biotech, Piscataway, NJ), and HDCBDysM was eluted with freezing buffer (50 mM HEPES, pH 7.5, 2 mM MgCl2, 150 mM NaCl, 5% sucrose). The viral titer was determined by measuring the optical density at 260 nm (25), and HDCBDysM DNA structure was confirmed by Southern blot analysis. The level of contamination by E1/E3-deleted helper adenovirus ranged between 0.02 and 0.17%, as determined by measuring the cytopathic effect after infection of 293A cells (28).
All aspects of the study were approved by the institutional animal ethics committee. Dystrophin-deficient mdx mice and wild-type C57BL10 mice were purchased from The Jackson Laboratory (Bar Harbor, ME). The mice (12 wk old) were anesthetized with ketamine (130 mg/kg) and xylazine (20 mg/kg) by intramuscular (im) injection. A laparotomy was performed to reveal the abdominal surface of the diaphragm. With the aid of a dissecting microscope, the mdx diaphragm was directly injected with AdV as previously described in detail (32). The mdx mice received injections of HDCBDysM (5 × 1010 vector particles, diluted in 100 μl of saline) into the hemidiaphragm on one side, whereas the contralateral side was injected with saline alone to serve as a within-animal control. To blunt any acute toxic inflammatory reaction to viral capsid proteins contained in the HDCBDysM inoculating dose (47), methylprednisolone was also given (2.5 mg·kg−1·day−1 im) for 3 days before and 2 days after vector administration. We have previously shown that this dose of methylprednisolone has no significant effects on the contractile properties of the mdx diaphragm (46). Animals recovered well from the surgery and demonstrated no untoward effects. The mice were killed at either 7 or 30 days after HDCBDysM administration.
Measurement of Diaphragmatic Contractility Parameters
Isometric contractile properties.
At 30 days postinjection, mice were anesthetized (130 mg/kg im ketamine and 20 mg/kg im xylazine) to achieve a loss of deep pain reflexes. HDCBDysM-treated and untreated mdx diaphragm strips were then carefully removed in random order to determine in vitro isometric contractile properties as previously described in detail (45). Briefly, the diaphragm strips were mounted vertically in a jacketed tissue bath chamber filled with Ringer solution continuously perfused with 95% O2:5% CO2 and maintained at 27°C (pH 7.4). The costal margin of each diaphragm strip was securely anchored to a platform near the base of the chamber while the central tendon was tied to the lever arm of a force transducer/length servomotor system (model 300B dual mode; Cambridge Technology, Watertown, MA). A mobile micrometer stage (Newport Instruments, Toronto, Canada) was employed to allow incremental adjustments of muscle length. Electrical field stimulation was induced via platinum plate electrodes placed into the bath on both sides of the muscle. Supramaximal stimuli with monophasic pulse duration of 2 ms were delivered using a computer-controlled electrical stimulator (model S44; Grass Instruments, Quincy, MA) connected in series to a power amplifier (model 6824A; Hewlett Packard, Palo Alto, CA). Muscle force was displayed on a storage oscilloscope (Tektronix, Beaverton, OR), and the data were simultaneously acquired to computer (Labdat/Anadat software; RHT-InfoData, Montreal, Quebec, Canada) via an analog-to-digital converter at a sampling rate of 1,000 Hz for later analysis. After adjusting each muscle strip to optimal length (Lo, the length at which maximal twitch force is achieved), we sequentially stimulated the muscles at low (10 Hz) and high (100 Hz) frequencies for 300 ms each, with a 2-min recovery period between each contraction; this allowed the spectrum of isometric tetanic force at the two ends of the force-frequency relationship to be ascertained. Diaphragm strips were then removed from the bath, and Lo was directly measured under a dissecting microscope with microcalipers accurate to 0.1 mm. Total muscle strip cross-sectional area was determined by dividing muscle weight by its length and tissue density (1.056 g/cm3). This allowed specific force (force/cross-sectional area) to be calculated, which was expressed as N/cm2.
Resistance to contraction-induced mechanical stress.
Muscle fibers lacking in dystrophin are abnormally susceptible to damage triggered by the high mechanical stresses associated with eccentric (i.e., lengthening) muscle contractions (34). Therefore, we subjected diaphragm strips to eccentric contractions and determined the resulting force deficit, which was defined as the percent decline in isometric force from the first to the last eccentric contraction. The latter served as a functional indicator of contraction-induced mechanical injury to dystrophic muscle as previously reported (7, 34). Each contraction involved supramaximal stimulation at 100 Hz for a total of 300 ms; the muscle was held at Lo during the initial 100 ms (isometric component) and then lengthened through a distance of 15% of Lo during the last 200 ms (eccentric component). Peak muscle length was maintained for an additional 100 ms after the cessation of electrical stimulation, followed by a return to Lo during the next 100 ms. A total of five such contractions were imposed on the muscle strip, each being separated by a 30-s recovery period. Last, a 100-Hz stimulation was performed at Lo to determine the final level of isometric force production at the end of the eccentric contraction protocol. Because the damage and isometric force deficit associated with eccentric contractions are directly correlated with the peak mechanical stress placed on the muscle (34), the force deficit was normalized to peak muscle stress as previously described (10, 11). All measurements were performed on HDCBDysM-treated and untreated diaphragm strips of mdx mice as well as in untreated diaphragm strips obtained from wild-type C57BL10 mice of the same age.
Immunohistochemistry and Morphometric Analysis
After completing the muscle function studies, we embedded diaphragms in mounting medium and snap-froze them in isopentane precooled with liquid nitrogen. Transverse sections (6-μm-thick) were obtained in a cryostat and then fixed on slides in 1% acetone.
Immunohistochemical procedures were carried out to detect dystrophin expression using a polyclonal antidystrophin (COOH terminus) primary antibody and biotinylated secondary antibody (45), with subsequent visualization by either horseradish peroxidase- or Cy3-conjugated streptavidin (Jackson ImmunoResearch Laboratories, West Grove, PA). Microscopically visualized sections were photographed using a digital camera, and the image was stored on computer. Analysis of the number and cross-sectional area of dystrophin-positive and dystrophin-negative myofibers on the entire diaphragm muscle strip cross section (averaging ∼800 myofibers) was then performed using image analysis software (ImagePro Plus; Media Cybernetics, Silver Spring, MD).
The cellular immune response to HDCBDysM injection of the mdx diaphragm was evaluated after 30 days by staining consecutive cryostat sections for CD4+ or CD8+ T lymphocytes as previously described (33). After anti-mouse CD4 (L3T4; BD Biosciences Pharmingen, Mississauga, Ontario, Canada) and CD8a (Ly2; Cedarlane, Hornby, Ontario, Canada) monoclonal antibodies raised in rat were applied, the sections were rinsed and reacted with appropriate biotinylated secondary antibodies (Jackson ImmunoResearch Laboratories). Specific staining was then visualized using the avidin-biotin peroxidase detection system (Vectastain ABC; Vector Labs, Burlingame, CA). For each muscle analyzed, computer-assisted image analysis (ImagePro Plus) was employed to determine the number of positively staining T lymphocytes in transduced areas. This was done by placing a rectangular grid encompassing ∼400 myofibers over the epicenter of the transduced area on the computer image; the number of CD4+ or CD8+ T lymphocytes was then determined and expressed per 100 myofibers (transduced as well as nontransduced) contained within the grid area.
All data are expressed as mean values ± SE. Data were analyzed using one- or two-way ANOVA, with post hoc application of Dunn's method where appropriate (SigmaStat; SPSS, Chicago, IL). Statistical significance was set at P < 0.05.
Stability of Dystrophin Expression in mdx Diaphragms After HDCBDysM Administration
After HDCBDysM injection of the mdx diaphragm, dystrophin-expressing fibers were found to be present in a somewhat nonuniform pattern of distribution throughout the muscle strip, as shown in Fig. 1. By immunohistochemistry, most dystrophin-expressing fibers demonstrated circumferential sarcolemmal staining for dystrophin at both 7 and 30 days post-HDCBDysM injection. In some instances, cytoplasmic staining was also observed, indicating that the level of dystrophin expression within these transduced fibers exceeded normal physiological levels. As expected, there was also a very small number of dystrophin-expressing fibers in the untreated mdx diaphragm, which are known to represent somatic cell mutations or so-called “revertant” fibers (17) that do not offer significant functional protection against the dystrophic process (9).
In HDCBDysM-treated mdx diaphragms, dystrophin expression was stable over the time course of the study. Hence, there was no significant change in the number of dystrophin-expressing fibers in the diaphragm between 7 and 30 days after HDCBDysM injection (see Fig. 2A). The relative contribution of dystrophin-expressing muscle fibers to total diaphragm muscle strip cross-sectional area was also determined (see Fig. 2B). This was found to comprise 20–25% of the total muscle area, and there was no significant alteration in the percentage of the diaphragmatic muscle area demonstrating dystrophin expression between the two time points examined.
Cellular Immune Response in mdx Diaphragms After HDCBDysM Administration
Transduced areas of the diaphragm injected 30 days earlier with HDCBDysM were immunostained for markers of CD8+ and CD4+ T lymphocytes, which play a central role in the immune-mediated loss of transgene expression after AdV-mediated gene transfer to muscle (33, 47). As shown in Figs. 3 and 4, there was a significant increase in the number of infiltrating T cells within the HDCBDysM-injected regions of the mdx diaphragm. There was a tendency for CD4+ T cells to outnumber the CD8+ T lymphocytes in the HDCBDysM-injected muscles, but this did not achieve statistical significance. It was hypothesized that the magnitude of the inflammatory response could be directly related to the number of transduced fibers if the latter represent an ongoing antigenic stimulus; alternatively, inflammatory cells could be responsible for the destruction of transduced fibers, in which case an inverse correlation would be expected. However, there was no significant correlation (either positive or inverse) between the number of dystrophin-expressing fibers in HDCBDysM-treated diaphragms and the levels of CD4+ or CD8+ lymphocytic infiltration.
Physiological Function of mdx Diaphragms After Full-Length Dystrophin Gene Transfer with HDCBDysM
At 1 mo after vector administration, there were no significant effects of HDCBDysM on either maximal twitch (2.9 ± 0.3 vs. 2.9 ± 0.4 N/cm2 during 1-Hz stimulation in treated and untreated, respectively), submaximal tetanic (4.6 ± 1.1 vs. 4.4 ± 0.5 N/cm2 during 10-Hz stimulation in treated and untreated, respectively), or maximal tetanic (10.1 ± 0.6 vs. 9.0 ± 0.9 N/cm2 during 100-Hz stimulation in treated and untreated, respectively) force production by the mdx diaphragm. Because a characteristic feature of dystrophin-deficient muscle is an abnormal susceptibility to injury caused by mechanical stress (34), we next determined the magnitude of the force deficits induced by imposing a series of eccentric contractions on the diaphragm. As shown in Fig. 5, there was a significant protective effect of HDCBDysM against eccentric contraction-induced force deficits. Hence, the force deficits induced by such high mechanical stress contractions were significantly lower in the HDCBDysM-treated mdx diaphragm strip compared with the untreated mdx muscle. It should be noted, however, that the protection afforded by HDCBDysM was not sufficient to completely normalize the response, as values in the HDCBDysM-treated mdx diaphragm remained intermediate between those seen in untreated mdx diaphragm strips and normal wild-type mice.
Different skeletal muscles do not necessarily respond identically to a lack of dystrophin, and this is equally true for potential therapeutic interventions for dystrophin deficiency. Indeed, we previously found that diaphragm and hindlimb (soleus) muscles of mdx mice exhibited different responses after being injected with first-generation AdV containing a 6.3-kb dystrophin minigene; the latter produced physiological improvements in the mdx hindlimb muscle but not in the diaphragm (45). Recently, gene expression profiling with oligonucleotide microarrays has revealed striking differences in the transcriptome between diaphragm and hindlimb muscles of normal as well as mdx mice (37). The majority of these differentially transcribed genes cannot be attributed to known fiber type differences. In fact, the differences between diaphragm and hindlimb (37) are substantially greater than between prototypical fast- and slow-twitch hindlimb muscles (1). Furthermore, the transcriptional pattern observed in diaphragm suggests a higher presence of antigen-presenting cells and other immune-responsive elements (37), suggesting at least one possible basis for the less favorable response to dystrophin gene transfer using first-generation AdV in the mdx diaphragm (45).
In the absence of dystrophin, muscle fibers are abnormally susceptible to contraction-induced tears of the muscle cell membrane, or sarcolemma (6, 34). Sarcolemmal damage may have several adverse consequences for muscle fibers, including depressed contractility and strength (34, 44), increased calcium influx into the muscle cells (24), and cellular necrosis. Because the mdx diaphragm is a physiologically relevant preclinical model for testing therapeutic interventions for DMD (43), we sought to evaluate the effects of full-length dystrophin gene transfer to the diaphragm in immunocompetent mdx mice using a fully gutted adenoviral vector. The major findings of our study were 1) full-length dystrophin gene transfer to the mdx diaphragm by HDCBDysM provided significant protection against the exaggerated loss of contractile force induced by high-stress muscle contractions in dystrophin-deficient muscles (34), and 2) despite mild T lymphocyte infiltration of the muscle, dystrophin expression in the mdx diaphragm was stable over a 30-day period without the need for continuous immunosuppressive therapy.
Physiological Effects of Gutted AdV in mdx Diaphragm
One of the most characteristic physiological findings in dystrophin-deficient muscles is an exaggerated loss of force-generating capacity after being subjected to high-stress eccentric contractions (7, 34). To our knowledge, this is the first report that dystrophin gene transfer with any viral vector can achieve functional benefits in the diaphragm with respect to this major hallmark of dystrophin deficiency. The magnitude of the effect was similar to that found after injection of the same vector into mdx hindlimb muscle (11). On the other hand, there was no significant difference in the nonstressed isometric force-generating capacity between untreated and AdV-treated diaphragms over the time course of our study.
Our findings are consistent with those recently reported by DelloRusso et al. (8) as well as Gilbert et al. (11) in adult mdx hindlimb muscles treated with gutted AdV encoding dystrophin. Therefore, our results and those of previous studies (7, 8, 11) indicate that a decreased vulnerability to the injurious effects of eccentric contractions is a sensitive marker of the early physiological efficacy of dystrophin gene replacement in mdx muscles. This is consistent with dystrophin's putative role as a cytoskeletal protein that provides mechanical reinforcement to the muscle cell but is not directly involved in the process of force generation. Hence, dystrophin replacement is not expected to acutely restore normal muscle strength but only to prevent or slow the process of further disease progression. It is likely that to demonstrate beneficial effects on isometric force production in muscles with established dystrophic pathology such as the mdx diaphragm, it would be necessary to observe the effects of treatment over a period of several months. Studies performed in transgenic mdx mice expressing dystrophin cDNAs have shown that dystrophin expression at ∼20% of wild-type levels is sufficient to prevent the myopathic phenotype (36). However, this differs from the situation in the current investigation, since in our study we treated mdx diaphragms postnatally at a time when substantial pathology and functional impairment were already present. In addition, it is possible that the percentage of transduced muscle fibers needed to ameliorate isometric force production under these conditions is greater than the level required to decrease force deficits caused by eccentric contractions.
We have previously demonstrated that in the absence of continuous immunosuppression, injection of first-generation AdV into the mdx diaphragm causes a profound decrease in the isometric force-generating capacity of the muscle (32). Under these conditions, both CD8+ and CD4+ T lymphocytes appear to play a role in the loss of force production by the diaphragm (33). In addition, leaky expression of viral gene products has a direct toxic effect on diaphragmatic muscle fibers, independent of cell-mediated immunity, which further compromises diaphragmatic contractility (33). Therefore, although dystrophin gene transfer with our gutted AdV did not improve isometric force-generating capacity of the diaphragm within the time frame examined in this study, it is important to note that adverse effects on isometric force previously observed after the use of first-generation AdV (32, 33) did not occur. This suggests that the margin of safety associated with treatment by gutted AdV is substantially greater than with first-generation vectors that still contain adenoviral genes.
The HDCBDysM vector employed in our study is deleted of all viral sequences except for the inverted terminal repeats and packaging signal (11). Because this vector cannot express viral gene products, both T cell-mediated destruction of transduced fibers and direct toxic effects on myofibers should be greatly mitigated. In addition, the increased cloning capacity of gutted AdV allows for use of the full-length dystrophin transgene, which is functionally superior to truncated forms of the gene used in other viral vectors (36). Indeed, the greatly increased insert capacity of our gutted AdV in particular (36 kb as opposed to ∼8 kb with earlier generation AdV) permitted delivery of not only one but two full-length dystrophin cDNAs per vector particle. In principle, this should allow one to achieve a higher level of therapeutic dystrophin gene delivery for a given inoculating dose, thereby further reducing any dose-dependent toxic or immune-mediated adverse effects on muscle function.
Cellular Immune Response to Gutted AdV in mdx Diaphragm
Without immunosuppressive therapy, transgene expression mediated by first-generation AdV is almost completely eliminated within 3–4 wk due to T cell-mediated destruction of the transduced cells (47, 48). In contrast, gutted AdV achieved sustained transgene expression in nondystrophic skeletal muscles without evidence of inflammation (2, 3). Several groups have reported similarly encouraging results with the use of gutted AdV in other models (20, 23, 26, 27, 39, 49). Although HDCBDysM was clearly superior to first-generation AdV (45) in its ability to avoid immune-mediated loss of transduced fibers in the mdx diaphragm, increased T lymphocyte infiltration of the diaphragm was nonetheless observed. Possible sources of this cellular immune response to HDCBDysM in the diaphragm include 1) the low-level contamination of the HDCBDysM preparation by nongutted helper virus (estimated as being 0.02–0.17%), which could not be completely eliminated with the available purification methods, 2) an immune reaction against murine dystrophin itself, since the latter may be treated as a neoantigen by the dystrophin-deficient host (12, 14, 29), and 3) a response to the viral capsid proteins contained in the initial AdV inoculum (19), even though the latter are more often associated with humoral rather than cellular adaptive immunity (47). Although T lymphocyte infiltration of the diaphragm was not associated with any reduction in dystrophin-expressing fibers or force-generating capacity in our study, previous work in adult mdx hindlimb muscles showed a gradual decrease in dystrophin expression several months after administration of HDCBDysM (11). Longer-term studies will be needed to determine whether this is also the case in the diaphragm.
In conclusion, we have found that full-length dystrophin gene transfer to the mdx diaphragm with a gutted adenoviral vector can achieve significant physiological reversal of a characteristic hallmark of dystrophic muscular dysfunction. In addition, stable dystrophin transgene expression was achieved in the mdx diaphragm over a 1-mo period, without apparent toxicity or a need for continuous immunosuppressive therapy. To the extent that the mdx diaphragm is a severely affected muscle that is phenotypically similar to human DMD, these findings suggest that gutted AdV may be a promising system for therapeutic gene transfer in this disease. However, a number of issues must be addressed before this can be considered. In particular, a more efficient delivery method is needed that will permit more widespread gene transfer within the diaphragm and other respiratory muscles. Although direct injection of the diaphragm could conceivably be performed via minimally invasive surgery (e.g., laparoscopy), it would be preferable to target a larger percentage of muscle fibers through the vascular system. This will likely be required to have a clinically significant positive impact on respiratory muscle strength. Recently, several investigators have made progress toward this goal (4, 13, 21). Finally, it will be important to identify the principal source of residual immune responsiveness to HDCBDysM or other gutted AdV and to determine whether a low but clinically acceptable level of immunosuppressive therapy is needed to sustain physiologically effective levels of dystrophin expression in the diaphragm over a period of many months or years.
This investigation was supported by grants from the Canadian Institutes for Health Research, the Muscular Dystrophy Associations of USA and Canada, and the Association Francaise Contre les Myopathies.
S. Matecki received a Postdoctoral Fellowship from the Canadian Institutes of Health Research. J. Nalbantoglu is a National Research Scholar of the Fonds de la recherche en sante du Quebec and a Killam Scholar. G. Karpati holds a Killam Chair of Neurology from the Montréal Neurological Institute. B. J. Petrof is a National Research Scholar of the Fonds de la recherche en sante du Quebec.
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