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Human Molecular Genetics, 2003, Vol. 12, No. 11 1287-1299
DOI: 10.1093/hmg/ddg141
© 2003 Oxford University Press

Prolonged dystrophin expression and functional correction of mdx mouse muscle following gene transfer with a helper-dependent (gutted) adenovirus-encoding murine dystrophin

Rénald Gilbert1,{dagger}, Roy W. R. Dudley2, An-Bang Liu1,{ddagger}, Basil J. Petrof2, Josephine Nalbantoglu1 and George Karpati1,*

1Neuromuscular Research Group, Montreal Neurological Institute, McGill University, Montréal, Québec, Canada H3A 2B4 and 2Respiratory Division, McGill University Health Center and Meakins-Christie Laboratories, McGill University, Montréal, Québec, Canada H3A 1A1

Received January 17, 2003; Accepted March 30, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Dystrophin gene transfer using helper-dependent adenoviruses (HDAd), which are deleted of all viral genes, is a promising option to treat muscles in Duchenne muscular dystrophy. We investigated the benefits of this approach by injecting the tibialis anterior (TA) muscle of neonatal and juvenile (4–6-week-old) dystrophin-deficient (mdx) mice with a fully deleted HDAd (HDCBDysM). This vector encoded two full-length murine dystrophin cDNAs regulated by the powerful cytomegalovirus enhancer/ß-actin promoter. At 10 days post-injection of neonatal muscles, 712 fibers (42% of the total number of TA fibers) were dystrophin-positive (dys+), a value that did not decrease for 6 months (the study duration). In treated juveniles, maximal transduction occurred at 30 days post-injection (414 dys+ fibers, 24% of the total number of TA fibers), but decreased by 51% after 6 months. All studied aspects of the pathology were improved in neonatally treated muscles: the percentage of dys+ fibers with centrally localized myonuclei remained low, localization of the dystrophin associated protein complex was restored at the plasma membrane, muscle hypertrophy was reduced, and maximal force-generating capacity and resistance to contraction-induced injuries were increased. The same pathological aspects were improved in the treated juveniles, except for reduction of muscle hypertrophy and maximal force-generating capacity. We demonstrated a strong humoral response against murine dystrophin in both animal groups, but mild inflammatory response occurred only in the treated juveniles. HDCBDysM is thus one of the most promising and efficient vectors for treating DMD by gene therapy.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Duchenne muscular dystrophy (DMD) is a fatal disease that affects one male birth in 3500 (1,2). It is caused by mutations of the dystrophin gene that encodes an elongated 427 kDa protein associated with the plasma membrane of skeletal and cardiac muscles. The hallmark of DMD is a progressive wasting and weakness of skeletal muscles leading to death around the age of 20. A promising approach to treat DMD is to restore dystrophin expression in the diseased muscle by injecting the muscle with naked DNA or viral vectors carrying the dystrophin cDNA. An ideal viral vector for treating DMD in this manner, in addition to being non-toxic, should be able to produce sufficient and sustained dystrophin expression in a major proportion of the body musculature.

Efficient and long-term transgene expression has been achieved in muscle after gene transfer using recombinant adeno-associated virus (rAAV) (3). One of the major problems with rAAV is its small insert capacity of about 5 kb, which is too small to carry the full-length dystrophin cDNAs (12 kb). Improvement of the histology and of the force-generating capacity of muscle was demonstrated after treatment of dystrophin-deficient (mdx) mouse muscle with rAAV carrying very small truncated version of dystrophin (microdystrophin) (4–7). However, it is unclear to what extent the expression of these microdystrophin cDNAs will mitigate the dystrophic phenotype in large animals and humans.

E1-deleted adenovirus (Ad), also known as first generation Ad (FGAd), can transduce muscle fibers in vivo. Indeed, several research groups have demonstrated the potential usefulness of FGAd-mediated dystrophin gene transfer (8–11), or that of its functional homolog utrophin (12–14), as a means of mitigating the dystrophic phenotype of the mdx mouse muscle. However, FGAd encode most of their viral genes. Low-grade synthesis from these genes after in vivo gene transfer triggers a cellular immune response that subsequently eliminates and/or silences transgene expression (15,16). In addition, the insert capacity of FGAd is only about 8 kb, which is sufficient for the Becker minidystrophin cDNA (6.5 kb), but not for the full-length dystrophin cDNA (12 kb). To reduce the immune response caused by FGAd, all the viral sequences except for the inverted terminal repeats (ITR) and the packaging signal of Ad were removed (17–20). Gene transfer using these completely deleted Ad vectors, also referred to as ‘gutted Ad’, large capacity Ad or helper-dependent Ad (HDAd), is associated with a reduction in the cellular immune response, which has led to an improvement in the duration of transgene expression (21–25). Another advantage of HDAd is the proportional gain in their insert capacity, which, in the case of a fully deleted HDAd, is increased to ~36 kb.

Despite their potential advantages as vectors for treating DMD by gene therapy, relatively few preclinical studies using HDAd-carrying dystrophin exist. Dystrophin gene transfer with HDAd in neonatal mdx mice can improve the muscle histology and can restore the dystrophin-associated protein complex (DPC) at the cell surface (26,27). Treatment of old mdx muscle with HDAd-carrying dystrophin can increase their resistance to contraction-induced injury but not their force production (28). We previously observed weak dystrophin expression level after treatment of mdx mouse and dystrophic golden retriever dog muscles with HDAd carrying dystrophin regulated by the cytomegalovirus (CMV) promoter (29). Because of this low expression level, we constructed an HDAd encoding two tandem human dystrophin cDNAs regulated by the powerful hybrid CMV enhancer/ß-actin (CB) promoter. However, despite excellent dystrophin expression level, the use of this vector still only resulted in transient dystrophin expression in mdx mouse muscle, most likely because dystrophin was non-isogenic (30). Transient dystrophin expression was also observed after treatment of mdx mouse using HDAd-encoding human dystrophin cDNA or murine dystrophin plus ß-galactosidase (26,31). Reduction of dystrophin expression was also observed even when transgenic mdx mice expressing ß-galactosidase were injected with the latter vector (27). However in this study, the mice were not completely tolerant to ß-galactosidase. Gene transfer of an isogenic dystrophin transgene using an HDAd into immunocompetent dystrophin-deficient muscle has thus not been completely investigated.

In the present study, to gain a better insight of the potential usefulness of HDAd for DMD treatment, we tested in mdx mouse muscle a fully deleted HDAd (HDCBDysM) encoding two tandem murine dystrophin full-length cDNAs, each regulated by the powerful (CB) promoter. Massive necrosis of the mdx mouse limb muscle fibers does not occur before 3–4 weeks of age (32). We thus treated the tibialis anterior (TA) muscle of neonatal and juvenile (4–6-week-old) mdx mice with HDCBDysM to verify if the pathology would be mitigated to the same extent when the muscle is treated before or after the onset of the necrotic process. In addition, treatment of neonatal and juvenile mdx muscle allowed us to compare the duration of dystrophin expression in animals with immature (neonatal) and fully developed immune system (juvenile).

Following injection of the TA muscle of neonatal mdx mice with HDCBDysM, we demonstrated unabated dystrophin expression to a level (more than 40% of the TA was transduced) and duration (6 months) not achieved before using an HDAd. In the treated juvenile mdx TA, maximal transduction occurred at 30 days post-injection (24% of the TA), but decreased by 51% after 6 months. An appreciable but mild inflammatory response, whose exact triggering agent is not known, was detected only in the juvenile-injected muscle. Marked improvement of muscle histology and physiology was achieved in both animal groups. HDCBDysM is thus a promising vector for treating DMD by gene therapy. Further studies are required to fully characterize the factor(s) triggering the inflammatory response, which might prevent long-term application of this vector in fully immunocompetent subjects.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Characterization of HDCBDysM
The structure of the fully deleted HDAd (HDCBDysM) encoding two identical murine dystrophin expression cassettes appears in Figure 1A. Dystrophin expression by this vector was controlled by the strong hybrid CMV enhancer/ß-actin (CB) promoter. We constructed this vector because efficient transgene expression using a similar HDAd encoding human dystrophin was demonstrated in muscle (30). HDCBDysM was produced using the 293Cre-loxP system (20) and purified by CsCl gradient centrifugation. To verify that the structure of HDCBDysM corresponded to the one depicted in Figure 1A, we isolated DNA from purified viral particles and analyzed it by Southern blot analysis. Following digestion with BamH1 and hybridization with a probe against the whole vector, or with a probe against the CB promoter, the observed bands pattern matched exactly the predicted structure of HDCBDysM (Fig. 1B).



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  		Figure 1. Characteristics of the vector used in this study. (A) Structure of HDCBDysM. The positions of the CB promoters, full-length murine dystrophin cDNAs, polyadenylation signals (pA), packaging signals ({Psi}), and left inverted terminals repeats of Ad (L-ITR) are indicated. The lines under HDCBDysM show the position of cleavage sites for BamHI. The numbers correspond to the size of the fragments in kilobases (kb) after digestion with BamHI. (B) Southern blot analysis of HDCBDysM DNA after digestion with BamHI. In lane a, the blot was hybridized to a probe consisting of the L-ITR, the dystrophin cDNA and CB promoter. In lane b, the blot was hybridized to a probe consisting of the CB promoter. The positions of DNA size markers (kb) is indicated at the left of the gel. A 0.6–0.7 kb band is observed in (lane a) after longer exposure.

 
Dystrophin expression in neonatally-injected muscle
To study the level and duration of dystrophin expression bestowed by HDCBDysM in vivo, we injected the TA of neonatal mdx mice with HDCBDysM at a titer of 2.0x1012 viral particles/ml and analyzed dystrophin expression by immunocytochemisty and by western blot at various time points. At 10 days post-injection, the earliest time point investigated, we observed an excellent transduction level, with an average of 712 fibers per muscle positive for dystrophin, a value corresponding to 42% of the fibers of the TA (Fig. 2A and B). Dystrophin expression was not homogeneous throughout the transduced area, as many fibers demonstrated an intense cytoplasmic immunostaining, a clear sign of dystrophin overexpression, whereas other fibers did not (see also Fig. 5A). Efficient dystrophin expression was further confirmed by western blot analysis, where the average quantity of dystrophin produced at 10 days post-injection in the tested muscles was 2.3 times the amount in wild-type mouse muscle (Fig. 2C). The number of dystrophin positive (dys+) fibers and the amount of dystrophin produced by the treated muscle, did not decrease at 30, 60, 90 and 180 days post-injection, indicating that the abundant early dystrophin expression was stable (Fig. 2B).



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  		Figure 2. Dystrophin expression after treatment of neonatal mdx muscle with HDCBDysM. The TA of neonatal mdx mice was injected with HDCBDysM at a titer of 2.0x1012 virus/ml and analyzed for dystrophin expression at the indicated time points. (A) Cryostat sections stained for dystrophin by immunhistochemistry at 10 (a), 30 (b), 60 (c), 90 (d) and 180 (e) days post-injection. Scale bar=250 µm, except for (a) where it is 125 µm. (B) Quantification of the number of dystrophin positive fibers. The data are the mean number of dystrophin positive fibers per TA±SEM. n, number of muscles analyzed. (C) Western blot analysis of dystrophin expression. 10 µg of two different muscle extracts for each time point post-injection in days (D) were analyzed using an antibody against dystrophin (DYS) or vinculin (VINC), which serves as loading control. Ten micrograms of normal mouse muscle (lane M) and non-injected mdx muscle (lane C) were also analyzed simultaneously.

 


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  		Figure 5. Dystrophin expression after treatment of juvenile mdx muscle with HDCBDysM. The TA of juvenile mdx mice was injected with HDCBDysM at a titer of 2.0x1012 virus/ml and analyzed for dystrophin expression at the indicated time points. (A) Cryostat sections stained for dystrophin by immunohistochemistry at 10 (a), 30 (b), 60 (c), 90 (d) and 180 (e) days post-injection. Scale bar=250 µm. (B) Quantification of the number of dystrophin positive fibers. The data are the mean number of dystrophin positive fibers per TA±SEM. n, number of muscles analyzed; the double asterisks mean significantly smaller than 10, 30 and 60 days; the single asterisk mean significantly smaller than 30 days. (C) Western blot analysis of dystrophin expression. Ten micrograms of two different muscle extracts for each time point post-injection in days (D) were analyzed using an antibody against dystrophin (DYS) or vinculin (VINC), which serves as loading control. Ten micrograms of normal mouse muscle (lane M) and non-injected mdx muscle (lane C) were also analyzed simultaneously.

 
Treatment of neonatal mdx mice with HDCBDysM mitigates the dystrophic phenotype
The mdx muscles are characterized by cycles of muscle fiber necrosis followed by regeneration. The regenerated fibers can be easily identified because they contain conspicuous internal myonuclei (33). To evaluate if expression of dystrophin by HDCBDysM could protect muscle fibers from necrosis, the percentage of fibers having centrally located nuclei (centronucleation index) was analyzed at various time points. For this study, we compared the centronucleation index of the dys+ fibers to the centronucleation index of the dystrophin negative (dys-) fibers of the same muscles (Fig. 3A). At 10 days post-injection, the centronucleation index was low (less than 7.5%) in both dys+ and dys- fibers and no significant difference existed between these two groups. At this early age, muscle fibers have not gone yet through cycles of necrosis and regeneration and have their nuclei still located at the periphery (32,33). At later time points, muscle necrosis occurred in the dys- fibers, which is reflected by their elevated centronucleation index (higher than 45%). In contrast, this index remained low in the dys+ fibers throughout the course of this study, indicating that treatment with HDCBDysM protects muscle from necrosis.



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  		Figure 3. Treatment with HDCBDysM protects mdx muscle from necrosis. The TA of neonatal (A) or juvenile (B) mdx mice were injected with HDCBDysM at a titer of 2.0x1012 virus/ml. At the indicated time points post-injection, the percentage of fibers with centrally located nuclei was determined in the dystrophin positive (Dys+) and dystrophin negative (Dys-) fibers of the injected muscles. The data are the mean number of dystrophin positive fibers per TA±SEM. n, number of muscles analyzed; the asterisks mean significantly less than dystrophin negative fibers.

 
The absence of dystrophin in muscle causes a dramatic secondary reduction of the DPC at the plasma membrane (34,35). To evaluate if dystrophin gene transfer would restore the normal distribution of the DPC at the cell surface, we stained cryostat sections of muscles that were treated 90 days earlier with HDCBDysM, for ß-dystroglycan and {alpha}-sarcoglycan, two major components of the DPC. In the control mdx muscle, the signal intensity of ß-dystroglycan and {alpha}-sarcoglycan at the cell surface was markedly reduced compared with wild-type mouse muscle. In contrast, the signal intensity of these two components in the dys+ fibers of the treated muscle was comparable to the signal found in wild type muscle, thus indicating restoration of the DPC at the cell surface (data not shown).

The lack of dystrophin in mdx muscle causes a reduction in force-generating capacity of the muscle, and increases its sensitivity to mechanical stress-induced injury following the application of lengthening (eccentric) contractions (36). To verify if treatment with HDCBDysM could improve the force-generating capacity of mdx muscle, we injected the right TA of neonatal mdx mice with HDCBDysM as described above, and the left TA, which was used a control, with only buffer. Isometric force parameters were then measured in both legs at 60 days post-injection and compared with age-matched wild-type control mice (Fig. 4A). Quantification of the transduction level of this experiment appears in Figure 2B. Maximal twitch force was significantly higher in HDCBDysM-treated mdx muscles relative to untreated TA muscles. In addition, the twitch force of the HDCBDysM-treated TA muscles was restored to normal wild-type levels. Maximal tetanic force was also greater in the HDCBDysM-treated mdx muscles compared with the contralateral untreated, but, in contrast to the twitch force data, remained lower than wild-type control mice.



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  		Figure 4. Treatment of neonatal mdx muscle with HDCBDysM improves muscle physiology. The right TA of neonatal mdx mice was injected with HDCBDysM (mdx-treated) and the contralateral TA with buffer (mdx-untreated). The force-generating capacity of the muscle was analyzed at 60 days post-injection. (A) Maximal isometric force production. (B) Schematic representation of physiological signals obtained during an eccentric contraction showing simultaneous tracings corresponding to muscle force, length change and stimulation. (C) Group mean data for the isometric force deficit induced by eccentric contractions are shown. (D) Representative signals from eccentric contractions imposed on C57 control, mdx-untreated and mdx-treated muscle. The solid line indicates the first eccentric contraction in a series of five such contractions, while the dashed line represents the tracing from the fifth eccentric contraction. In the C57 control and treated mdx muscles, note the relative maintenance of isometric force at the fifth eccentric contraction in comparison to the untreated mdx. Values are group means±standard error. n, number of muscles analyzed.

 
The increased susceptibility of dystrophin-deficient myofibers to damage by eccentric contractions is manifested by a significant reduction in maximal isometric force production after each eccentric contraction. In addition, the level of damage and dysfunction induced by eccentric contractions is directly correlated with the magnitude of peak mechanical stress imposed on the muscle (36). Therefore, the effect of eccentric contractions on maximal isometric force production by the TA was determined, and this was normalized to account for any differences in peak mechanical stress placed on the muscles. As shown in Figure 4C and D, the isometric force deficit after eccentric contractions was significantly lower in HDCBDysM-treated mdx muscles than in the contralateral buffer-injected TA. However, the force deficit after eccentric contractions remained higher in HDCBDysM-treated mdx muscles compared with wild-type control mice. Inflammation caused by intramuscular injection of FGAd encoding an immunogenic transgene such as ß-galactosidase resulted in the upregulation of extrasynaptic utrophin, a functional dystrophin homolog (37). To verify that treatment with HDCBDysM did not upregulate utrophin in our study, sections of muscles treated 60 days earlier with HDCBDysM were immunostained for utrophin. No difference of the extrasynaptic utrophin staining intensity was observed between the muscle injected with HDCBDysM and the muscle injected with the buffer (data not shown). Thus, treatment with HDCBDysM does not upregulate extrasynaptic utrophin expression.

The limb muscles of mdx mice are hypertrophic and weigh significantly more than normal muscle (32). To determine if treatment with HDCBDysM would reduce the hypertrophy, the weight of the TA treated 60 days earlier with HDCBDysM was compared to contralateral buffer-injected TA. The average muscle weight of the TA injected with HDCBDysM was 47.5±1.7 mg (n=17) which was significantly less (P<0.001) than the average of 63.4±2.2 mg observed for the contralateral control TA (n=17). This result indicated that treatment with HDCBDysM at an early age reduces muscle hypertrophy.

Prolonged dystrophin expression after treatment of juvenile mdx mice with HDCBDysM
Neonatal mice can become partly tolerant to the Ad vector and transgene because they do not have a completely matured immune system. For this reason, and to verify the extent to which treatment of HDCBDysM could mitigate the pathology after the onset of muscle necrosis, we repeated all of the above experiments using juvenile (4–6-week-old) mdx mice, which were fully immunocompetent. We injected the TA of juvenile mdx mice with HDCBDysM at a titer of 2.0x1012 viral particles/ml and analyzed dystrophin expression by immunocytochemistry and by western blot analysis at various time points. At 10 days post-injection, the earliest time point investigated, a good transduction level was observed because an average of 276 dys+ fibers were counted in the treated TA, a value corresponding to 16% of the total fibers of the TA (Fig. 5A and B). The average number of dys+ fibers was 415 (24% of the TA) at 30 days post-injection and remained high at 60 days post-injection. However, a slight but significant reduction in the number of dys+ fibers occurred at 90 and 180 days post-injection. Despite this, an average of 204 muscle fibers were still dys+ at 180 days post-injection. In previous experiments, we never observed such a prolonged transgene expression following gene transfer in juvenile mdx muscle using FGAd encoding dystrophin or utrophin, or using a fully deleted HDAd encoding human dystrophin (12,13,30,38,39). The presence of full-length dystrophin in the treated muscle was also demonstrated by western blot analysis at all the time points investigated (Fig. 5C). The highest amount of dystrophin was observed in the muscle analyzed at 30 and 60 days post-injection. At these two time points, the average quantity of dystrophin produced in the four muscles analyzed corresponded to 33% of the amount of wild-type murine muscle.

HDCBDysM treatment of juvenile mdx muscle mitigates the dystrophic phenotype
We computed the centronucleation index of the dys+ fibers of juvenile-injected muscles at various time points and compared this value to the index of dys- fibers of the same muscle. At every time point investigated, the centronucleation index was lower in the dys+ fibers, thus demonstrating a protective effect against necrosis (Fig. 3B). The centronucleation index was much higher in the dys+ fibers of juvenile-treated muscles compared with the neonatally treated ones, because at the time of injection, a significant proportion of muscle fibers already had centrally located nuclei (Fig. 3A). The centronucleation index of the dys+ fibers did not decrease significantly at later time points, indicating the internally located nuclei do not readily move back to the periphery. We also stained for ß-dystroglycan and {alpha}-sarcoglycan, cryostat sections of juvenile-injected mdx muscles that were treated 60 days earlier with HDCBDysM. The signal intensity of these two components was increased at the cell surface of the dys+ fibers of the treated muscle, thus indicating restoration of the DPC at the plasma membrane (data not shown).

We also tested if HDCBDysM would ameliorate the physiological indices of the juvenile-treated muscle. For this experiment, in an attempt to increase the transduction level, the right TA of juvenile mdx mice was injected twice (78 h between each injection) with HDCBDysM as described above and the left TA was injected with buffer. To avoid potential adverse effects on transduction efficiency caused by greater non-antigenic-specific inflammation (innate immune response) triggered by the second exposure to HDCBDysM capsid proteins (40), mice were transiently immunosuppressed with FK506 (38) for 7 days, starting one day before treatment with HDCBDysM. Isometric force parameters were then measured in both legs at 60 days post-injection and compared with age-matched wild-type control mice. In contrast to the observation made with neonatally injected muscle, no improvement of the maximal twitch force and maximal tetanic force was observed (Fig. 6A). The lack of improvement was not due to poor transduction level, because the average number of dys+ in the six muscles used for the physiology was 345±102. This value was not statistically different from the one obtained after a single injection of HDCBDysM (Fig. 5B). The effect of eccentric contractions on maximal isometric force production by the TA was also determined. As shown in Figure 6B, the isometric force deficit after eccentric contractions was significantly lower in HDCBDysM-treated mdx muscles than in the contralateral buffer-injected TA, but remained, nonetheless, higher than wild-type control mice.



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  		Figure 6. HDCBDysM treatment of juvenile mdx muscle improves resistance to contraction-induced injury. The right TA of juvenile mdx mice was injected with HDCBDysM (mdx-treated) and the contralateral TA with buffer (mdx-untreated). The force-generating capacity of the muscle was analyzed at 60 days post-injection. (A) Maximal isometric force production. (B) Group mean data for the isometric force deficit induced by eccentric contractions are shown. C57 control, TA of normal mice of the same age. Values are group means±standard error; n, number of muscles analyzed.

 
The weight of the TA treated 60 days earlier with two doses of HDCBDysM was compared with the contralateral TA. In contrast to the observation made with the neonatally injected muscles, the average muscle mass of the juvenile TA treated with HDCBDysM (59±5 mg, n=6) was not significantly different from the control TA (59±3 mg, n=6).

Immune response after treatment with HDCBDysM
To test if injection of HDCBDysM would trigger a cellular inflammatory response, the transduced areas of muscle injected once 60 days earlier with HDCBDysM were stained for markers for macrophages, CD8+ and CD4+ T-lymphocytes. No significant increase of immune cells compared with mdx control muscles occurred in the neonatally injected animals (Fig. 7). In contrast, we observed slightly more CD4+ and CD8+ T-lymphocytes in the transduced areas of the juvenile-injected animals, but the number of macrophages was not higher. Thus, treatment with HDCBDysM causes a slightly stronger inflammatory response in juvenile-injected compared to neonatally-injected muscles in agreement with a previous study (27). However, we considered this inflammatory response to be mild, because significantly more immune cells were observed after treatment with a FGAd encoding dystrophin (Fig. 7A). This mild inflammatory response was clearly insufficient to eliminate all the dys+ fibers, because an average of 204 fibers per TA were still expressing dystrophin at 180 days after injection of HDCBDysM (Fig. 5B). Because mdx muscles do not express full-length dystrophin, we then determined if treatment with HDCBDysM would generate a humoral response against dystrophin. Sera were collected from the treated animals and analyzed for the presence of antibody against murine dystrophin by western blot analysis using lysate of cells transfected with the murine dystrophin cDNA as a source of antigen. All the sera analyzed (seven juvenile-injected and six neonatally injected animals) demonstrated the presence of antibody against the murine dystrophin (data not shown).



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  		Figure 7. Treatment of juvenile muscle with HDCBDysM generates a mild inflammatory response. (A) The TA of juvenile (Adult) and neonatal (Neo) mdx mice were injected with HDCBDysM at a titer of 2.0x1012 virus/ml. Sixty days later, cryostat sections of injected muscles were stained using antibody against macrophages (Mac), CD4 or CD8 T-lymphocytes. Buffer-injected juvenile mdx TA (Cont) at 60 days-post-injection, and juvenile mdx TA injected with FGAd encoding the human minidystrophin cDNA (FGAd) (10) at 10 days post-injection, were also analyzed. The data are the means±SEM; n, number of muscle analyzed. The asterisks mean significantly higher than buffer-injected mdx; the hashes mean significantly smaller than FGAd-injected mdx. (B) Example of mild inflammatory response in juvenile-treated muscle. Consecutive cryostat sections of mdx TA injected 60 days earlier with HDCBDysM were stained for dystrophin (a), macrophages (b), CD4 (c) and CD8 T-lymphocytes (d). Arrows: localization of inflammatory cells in the muscles. Scale bar=210 µm.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
The present work represents the most comprehensive published study of full-length isogenic dystrophin gene transfer into mdx muscle using an HDAd in both neonatal and older animals. To obtain high-level dystrophin expression in vivo, we employed a unique and efficient Ad vector (HDCBDysM) encoding double tandem dystrophin expression cassettes controlled by a very strong hybrid promoter. No extraneous ‘stuffer’ sequences were present in this vector: it contained only the expression cassette flanked by the Ad ITR, and the Ad packaging signal. We used neonatal and juvenile mdx mice to verify that the pathology would be mitigated to the same extent when the muscle is treated before or after the onset of necrosis, and to compare dystrophin expression in animals with immature and fully developed immune system. We believe juvenile mdx mice are a better model to investigate the immune response on dystrophin expression, compared with old mdx mice (1-year-old) used in a recent study (28), because of higher prevalence of immune cells in juvenile mdx muscles (2). Our major findings are unabated dystrophin expression, and correction of all pathological aspects investigated in neonatal animal, despite induction of a strong humoral response again the murine dystrophin. In contrast, in juvenile-treated muscle, a slight but significant reduction of dystrophin expression occurred at 90 and 180 days post injection, and muscle hypertrophy as well as maximal isometric force-generating capacity were not ameliorated. Our data thus suggests that treatment of dystrophin-deficient muscle by gene therapy before the onset of necrosis would mitigate the pathology more efficiently, and that cellular immune response might prevent long-term (more than 6 months) application of our vector in fully immunocompetent individuals.

A successful treatment of DMD using a dystrophin gene transfer approach will most likely require that a large proportion of the body muscle mass is transduced and expresses significant and sustained amount of dystrophin. In an attempt to achieve this goal, we have previously constructed a fully deleted HDAd encoding two human dystrophin cDNAs controlled by the strong CB promoter (30). Although efficient, dystrophin expression was transient in mdx muscle, most likely because the dystrophin used was of human origin. In the present study, using the identical vector, but this time carrying the murine (isogenic) dystrophin cDNA, we demonstrated efficient and prolonged dystrophin expression after gene transfer in mdx mouse muscle. The fact that expression of murine dystrophin transgene was more stable compared with human dystrophin transgene is in agreement with various studies (41–43), demonstrating that isogenic proteins (murine dystrophin) are expressed for longer periods of time in muscle.

In juvenile mdx muscles that were treated with HDCBDysM, we observed a mild cellular infiltration by T-lymphocytes. The presence of low level of immune cell infiltrates was also observed by other researchers after gene transfer of fully deleted HDAd carrying the murine dystrophin cDNA in adult mdx muscle (27,28). It is possible that the reduction of transgene expression we observed at 90 and 180 days post-injection was due to destruction of some of the transduced fibers by the immune cell infiltrate (41,44) and/or resulted from down-regulation of the transgene expression due to the release of cytokines by the immune cells (45,46). The exact nature of the immunological insult responsible for triggering this low level of inflammation in the muscle is not known. The muscles of mdx mice do not normally synthesized the full-length dystrophin and, thus, the recombinant murine dystrophin could appear as a neoantigen and trigger the immune response. This is supported by the fact that dystrophin antigen produced by transplanted myoblasts in dystrophin-deficient mice can induce a cytotoxic T-lymphocyte response specific for dystrophin (47). We and others (27) have demonstrated the presence of a humoral response against murine dystrophin in the mdx mice treated with HDAd encoding dystrophin cDNA, indicating it could act as a neoantigen. In contrast, no humoral response was observed after intramuscular injection of plasmid DNA encoding murine dystrophin in mdx mice (48). The reason for this discrepancy is not clear. The Ad viral capsids could act as an immunological adjuvant. Alternatively, the immune response against murine dystrophin could be due to the higher transduction level conferred by the HDAd. However, it is unlikely the humoral response against dystrophin was responsible for the loss of transgene expression, because no such loss occurred in neonatally injected animals, despite the fact that a comparable humoral response against dystrophin was generated. Normal and dystrophin deficient muscles contain low levels of utrophin, a functional analog of dystrophin (49). Forced utrophin expression in transgenic animals or after gene transfer using FGAd (12,13,50,51) can mitigate the pathology associated with the lack of dystrophin. Utrophin gene transfer is thus an interesting alternative for treating DMD, because this protein is already present in dystrophin-deficient muscle and its overexpression should be well tolerated by the immune system of the host. Another approach to reducing the immune response against the transgene would be to control its expression using a muscle specific promoter (44,52). However, a clear advantage of this strategy to treat DMD using HDAd technology has yet to be demonstrated (27). Another factor that could trigger the inflammatory response is the low-grade synthesis of viral protein from the residual level of helper virus contaminating our preparation of HDAd. Long-term transgene expression has been achieved in healthy adult muscle after gene transfer with FGAd and with HDAd encoding isogenic transgene and containing comparable level of helper Ad (23,24,41,42). The mdx muscle, because of its chronic inflammatory state (due mostly to phagocytes) (44,53,54), is presumably highly sensitive to the presence of non-self proteins, and for this reason, even low level of helper Ad contamination could be sufficient to trigger a damaging immune response.

The excellent and prolonged transduction level obtained after treatment of neonatal and juvenile mdx mice with HDCBDysM allowed us to compare the extent to which this treatment mitigates the pathology when the muscle is treated before (neonates) or after the onset of the necrotic process ( juveniles). After treatment of neonates with HDCBDysM, the centronucleation index remained low in the dys+ fibers, indicating the transduced fibers were protected from necrosis for the complete duration of the study. In addition, the localization of the DPC was restored at the plasma membrane and we observed a significant increase in the resistance to contraction-induced injury as well as an improvement of isometric force generating capacity of the muscle. Comparable reduction of the centronucleation index and restoration of the DPC at the cell surface were documented following gene transfer of neonatal mdx mice using HDAd-encoding dystrophin (26,27) or with FGAd carrying dystrophin or utrophin (8,10,12,55). Our current study is the first to analyze the muscle physiology after treatment of neonates with an HDAd. In this respect, our data are comparable to those obtained with FGAd carrying dystrophin or utrophin in neonates (9,12,13). We further demonstrated that treatment with HDCBDysM is able to significantly reduce muscle hypertrophy, suggesting that homeostasis of muscle fiber volume is abnormal in the absence of dystrophin. Although no previous dystrophin gene transfer study exists about improvement of muscle hypertrophy in neonates, transgenic mouse experiments have shown that forced dystrophin or utrophin transgene expression in mdx muscle can reduce muscle weight (7,56).

At the time of injection, about 50% of the fibers of juvenile mdx TA have gone through at least one cycle of muscle necrosis followed by regeneration, and consequently possess centrally located myonuclei (32). The fact that we were able to demonstrate a stable 35% reduction of the centronucleation index after treatment of juvenile mdx muscle with HDCBDysM indicated that muscle fibers having peripheral nuclei at the time of transduction were subsequently protected from necrosis. Improvement of the centronucleation index of juvenile and adult mdx muscle was also documented after treatment with FGAd carrying utrophin or dystrophin transgenes (11,13), or after treatment with rAAV carrying microdystrophin (4,7). In one study (4), in agreement with our data, the percentage of dys+ fibers with centrally located nuclei did not decrease at later time points, indicating the internally located nuclei do not readily move to the periphery. In contrast, a significant reduction of the percentage of dys+ fibers with centrally located nuclei was observed at 5 months post-injection in another study (7). The reason for this apparent discrepancy is not clear. Because we observed a mild inflammatory response in the juvenile mdx muscles treated with HDCBDysM, we cannot rule out the presence of cellular infiltrate being responsible for maintaining the myonuclei in a central position. In juvenile muscles treated with HDCBDysM, we observed restoration of the amount of the DPC at the cell surface and a significant increase of the resistance to contraction-induced injury, which, however, still remained lower than normal muscle. By contrast, we did not observe an increase of the normalized specific isometric force. Similar observations concerning these two physiological parameters were recently made following dystrophin gene transfer in adult mdx muscle using HDAd (28). In contrast, improvement of the specific muscle isometric force was demonstrated after gene transfer of microdystrophin in adult mdx muscle using AAV (5). In the same study, an improvement of the resistance to contraction-induced injury was also demonstrated, which, however, was not statistically different from that of normal muscle. It is not clear why these parameters of functional muscle recovery were better after gene transfer with AAV, despite the fact the transgene was a microdystrophin. However, it is worth mentioning that the transduction efficiency was slightly higher with AAV, and that treatment with the latter vector did not increase the number of cytotoxic T-lymphocytes in the transduced muscle areas (4,5), when compared with HDAd (28). Finally, in agreement with our data, treatment with AAV (5) and HDAd (28) of adult mdx muscle did not reduce muscle hypertrophy.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Construction, purification and characterization of HDCBDysM
To construct a HDAd encoding the full-length cDNA of murine dystrophin regulated by the strong hybrid CB promoter, we first made plasmid pCBmDys5'ITR. The muscle creatine kinase promoter of pMDA which also contains the full-length murine dystrophin cDNA (57) was removed by digestion with XhoI and replaced with the CB promoter obtained by digesting pCAGGS (58) with XhoI and SalI. The resulting plasmid was further modified by inserting into the SpeI/NotI sites at the 3' end of dystrophin, the ß globin poly (A) signal followed by the packaging signal and the 5' inverted terminal repeat (ITR) of Ad type V derived from pAdCMV-dys (10). Plasmid pCBmDys5'ITR thus contains (in a 5' to 3' orientation) a unique NotI site, the 5'-ITR and the packaging sequence of Ad, the rabbit ß-globin poly (A) signal, the full-length murine dystrophin cDNA, the CB promoter and a unique XhoI site. HDCBDysM was generated by digesting pCBmDys5'ITR with NotI and the ends were dephosphorylated. After phenol–chloroform extraction and ethanol precipitation, the DNA was digested with XhoI, and the NotI–XhoI fragment containing the dystrophin expression cassette was purified by electrophoresis. The purified NotI–XhoI fragment was self-ligated, extracted by phenol–chloroform, and precipitated by ethanol. The ligation product was used to transfect 293Cre4 cells (20) with LipofectAMINETM (Life Technologies, Burlington, Ontario, Canada) according to the manufacturer's recommendations. HDCBDysM was then amplified using 293Cre–loxP system and purified by two consecutive continuous CsCl gradient centrifugations as described previously (29). After ultracentrifugation, the CsCl was removed by chromatography on Sephadex G25 columns (Amersham Pharmacia Biotech Inc., Piscataway, NJ, USA) and HDCBDysM was eluted with 50 mM HEPES pH 7.5, 2 mM MgCl2, 150 mM NaCl, 5% sucrose (freezing buffer). The titer (virus particles/ml) was determined by measuring the optical density at 260 nm (59) and the level of helper virus contamination, which was determined by measuring the cytopathic effect after serial dilution on 293A cells (60) ranged between 0.02 and 0.17% in different vector batches. When not used the same day, the vector was kept at -80°C. We confirmed the structure of the viral DNA by restriction analysis of purified DNA (61) followed by Southern blot as described previously (30). The probe consisted of a purified 16 kb NotI/XhoI DNA fragment of pCBmDys5'-ITR containing the 5'-ITR and the complete dystrophin expression cassette, or a purified 1.6 kb SalI–XhoI DNA fragment of pCCAGS containing the CB promoter.

Animal injection
We performed all animal experiments according to McGill University guidelines for animal care. We injected the left and right TA muscles of neonatal (2–4-day-old) or juvenile (4–6-week-old) mdx mice (C57BL/10ScSn-mdx/J; The Jackson Laboratory, Bar Harbor, ME, USA) once with 10 or 30 µl, respectively, of Ad vectors at a titer of 2x1012 virus particles/ml as described previously (10). Before the injection, the juvenile mdx mice were anesthetized by intraperitoneal injection of 2.5% Avertin. At 10, 30, 60, 90 or 180 days post-injection, the mice were euthanized by an overdose of pentobarbital, the TA was removed and frozen in liquid nitrogen-cooled isopentane.

Staining and western blot of muscle tissues
Transverse cryostat sections were stained for dystrophin, using a rabbit polyclonal antibody raised against the C-terminus of human dystrophin that recognizes murine and human dystrophin (10). We visualized the staining using either horseradish peroxidase- or Cy-3-conjugated streptavidin (Jackson Immuno Research Laboratories Inc., West Grove, PA, USA). For each injected muscle, we determined the total number of transduced fibers by counting the number of dys+ fibers on a single cryostat section, which spans the entire TA cross-section. To calculate the percentage of transduced fibers, the number of dys+ fibers was divided by 1700, which corresponds to the mean number of fibers in the TA muscle of 1–9-week-old mdx mice. We calculated the percentage of muscle fibers with centrally located nuclei by counterstaining with hematoxilin cryostat sections that were previously stained for dystrophin. Some sections were also stained using antibody against ß-dystroglycan (NCL-43DAG; Novocastra, Newcastle-upon-Tyne, UK), {alpha}-sarcoglycan (NCL-50DAG; Novocastra) and utrophin (NCL-DRP2; Novocastra), as described previously (55). Dystrophin expression was analyzed by western blot using 10 µg of muscle protein (30). The membrane was processed using the ECF western blotting kit (Amersham Pharmacia Biotech, Buckinghamshire, UK) according to the manufacturer's recommendations. The signal was visualized and quantified using a PhosphorImager system (STORM, Molecular Dynamics Inc., Sunnyvale, CA, USA). The blots were also stained with a monoclonal antibody against vinculin (V2638, Sigma-Aldrich, St Louis, MO, USA) as a loading control.

Cellular and humoral response
We measured the cellular immune response caused by HDCBDysM by staining consecutive cryostat sections for dystrophin as described above, and for macrophages, CD8+, or CD4+ T-lymphocytes using rat monoclonal antibodies Mac1, anti-CD8a (Ly 2, Cedarlane, Hornby, Ontario, Canada) and anti-CD4 (L3T4, BD Biosciences Pharmingen, Mississauga, Ontario, Canada), respectively. Mac1 was prepared from supernatant of rat hybridoma (M1/70.15.11.5HL) obtained from the American Type Culture Collection (Rockville, MD, USA). The signal was visualized using a rat anti-mouse biotinylated antibody (Jackson Immuno Research Laboratories Inc., West Grove, PA, USA) followed by horseradish peroxidase-conjugated streptavidin (Jackson Immuno Research Laboratories Inc.). For each muscle analyzed, we computed the number of positive inflammatory cells in the transduced areas (regions with more than 50% dys+ fibers), and we divided this value by the total number of fibers (dys+ and dys-) of the area. The TA of age-matched mdx mice previously injected with freezing buffer, and the TA of juvenile mdx injected with a FGAd encoding human dystrophin (AdCMV-dys) (10) at a titer of 1.4x1012 virus particle/ml and analyzed 10 days later, were used as controls. We measured the humoral response against murine dystrophin by transfecting dishes of 293A cells with pCBmdys5'ITR using Cytofectene (Bio-Rad Laboratories, Hercules, CA, USA) according to the manufacture's recommendations. The next day, the cells were lysed and 50 µg of protein were separated on a 5% gel and processed for western blot by enhanced chemiluminescence as described previously (29). The sera of uninjected mdx mice or of mdx mice injected with HDCBDysM were used a primary antibody. The detection of a dystrophin band on the blot indicated the presence of antibodies against dystrophin in the tested serum.

In vivo measurement of force generation
To measure the force of neonatally-injected muscle, the right TA of 2–4-day-old mdx mice were injected as described above with HDCBDysM. The left TA, which was used as control, was injected with 10 µl of freezing buffer. To measure the force of juvenile-injected muscle, 4–6-week-old mdx mice were immunosupressed by daily subcutaneous injections of FK506 (a generous gift of Fujisawa Inc., Japan) for 7 days. On the second and fifth day of FK506 treatment, the right TA was injected with 30 µl of HDCBDysM as described above and the left TA with 30 µl of freezing buffer. At 60 days post-injection, mice were anesthetized with ketamine (130 mg/kg) and xylazine (20 mg/kg) to achieve a loss of deep pain reflexes and immobilized in the supine position. Two 27.5 gage needles were used to secure the knee and ankle to a surgical platform. The distal tendon of the TA muscle was isolated and tied with 4-0 nylon suture to the lever arm of a force transducer/length servomotor system (model 305B dual mode; Cambridge Technology, Watertown, MA, USA). The latter was mounted on a mobile micrometer stage to allow fine incremental adjustments of muscle length. Exposed portions of the TA were kept moist with a 37°C isotonic saline drip, and the TA was then stimulated directly via an electrode placed on the belly of the muscle. Supramaximal stimuli with a monophasic pulse duration of 2 ms were delivered using a computer-controlled electrical stimulator (model S44; Grass Instruments, Quincy, MA, USA). Muscle force and length signals were displayed on a storage oscilloscope (Tektronix, Beaverton, OR, USA) and simultaneously acquired to a computer (Labdat/Anadat software; RHT-InfoData, Montreal, Canada) via an analog- to-digital converter at a sampling rate of 1000 Hz.

After adjusting the TA to optimal muscle length (L0, the length at which maximal twitch force is achieved), two twitch stimulations were recorded and the mean value was considered as maximal isometric twitch force. Maximal isometric tetanic force was then measured by stimulating the muscle at 120 Hz for 300 ms, allowing a clear plateau in force to be attained. Following a 2 min recovery period, the ability of the TA to withstand a series of high-stress eccentric (lengthening) contractions was determined. Each contraction involved supramaximal stimulation at 120 Hz for a total of 300 ms; the muscle was held at L0 during the initial 100 ms (isometric component), and then lengthened through a distance of 25% of L0 during the last 200 ms (eccentric component). Peak muscle length was maintained for an additional 100 ms after cessation of the stimulation, followed by a return to L0 during the next 100 ms. A total of five such contractions were imposed on the TA, each being separated by a 2 min recovery period. Lastly, a 120 Hz stimulation was performed at L0 to determine the final level of isometric force production following the eccentric contraction protocol. The isometric force deficit induced by each eccentric contraction was normalized to take into account the magnitude of mechanical stress placed upon the muscle. This was done by dividing the percent force drop by the level peak stress (N/cm2) attained during the preceding eccentric contraction. All measurements were performed in HDCBDysM-treated and untreated (contralateral limb) TA muscles of mdx mice, as well as in untreated TA muscles of normal wild-type C57Bl/10 mice (The Jackson Laboratory, Bar Harbor, ME, USA) or normal wild type C57Bl/6 mice (Charles River Laboratory, St-Constant, Quebec, Canada) of the same age. We did not observe any differences between the force generating capacity of C57Bl/10 and C57Bl/6.

Statistical analysis
Unless stated otherwise the data are expressed as the mean±SEM. The data were analyzed using an unpaired two-tailed t-test, or with an analysis of variance (ANOVA) followed by the Fisher's LSD procedure to compare the means. Comparisons of the response to eccentric contractions were analyzed by two-way ANOVA, with both treatment status and the number of eccentric contractions being incorporated into the ANOVA model. Statistical significance was set at P<0.05.


   ACKNOWLEDGEMENTS
 
We thank Jun-Ichi Miyazaki (Osaka University Medical School, Osaka, Japan) for the generous gift of plasmid pCAGGS, as well as Merk and Co., Inc. (West Point, PA, USA) for the 293Cre4 cells and AdLC8cluc. We acknowledge the expert technical assistance of Carol Allen, Mylène Bourget, Stephen Prescott and Klara Rostworowski. J.N. is a National Scholar of the Fonds de la recherché en santé du Québec and a Killam Scholar. This work was supported by the Canadian Institutes for Health Research, the Muscular Dystrophy Association of USA and Canada, and the Association Francaise Contre les Myopathies. This is an NRC publication number 37701.


   FOOTNOTES
 
* To whom correspondence should be addressed at: Montreal Neurological Institute, 3801 University Street, Montréal, Québec, Canada H3A 2B4. Tel: +1 5143988528; Fax: +1 5143988310; Email: george.karpati{at}mcgill.ca Back

{dagger} Present address: Biotechnology Research Institute, NRC, 6100 Royalmount Ave, Montréal, Québec, Canada H4P 2R2. Back

{ddagger} Present address: Department of Neurology, Tzu Chi Medical Center, Hualien, Taiwan. Back


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 DISCUSSION
 MATERIALS AND METHODS
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