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Human Molecular Genetics Advance Access originally published online on April 4, 2006
Human Molecular Genetics 2006 15(10):1610-1622; doi:10.1093/hmg/ddl082
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© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

A highly functional mini-dystrophin/GFP fusion gene for cell and gene therapy studies of Duchenne muscular dystrophy

Sheng Li1,2, En Kimura1,2, Rainer Ng3, Brent M. Fall1,2, Leonard Meuse1,2, Morayma Reyes1,2, John A. Faulkner3,4 and Jeffrey S. Chamberlain1,2,*

1Department of Neurology and 2Senator Paul D. Wellstone Muscular Dystrophy Cooperative Research Center, University of Washington School of Medicine, Seattle, WA 98195-7720, USA and 3Department of Biomedical Engineering and 4Department of Molecular and Integrative Physiology, University of Michigan, Ann Arbor, MI 48109-2007, USA

* To whom correspondence should be addressed at: K243b HSB, University of Washington School of Medicine, 1959 NE Pacific Street, Seattle, WA 98195-7720, USA. Fax: +1 2066168272; Email: jsc5{at}u.washington.edu

Received February 16, 2006; Accepted March 25, 2006


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
A promising approach for treating Duchenne muscular dystrophy (DMD) is by autologous cell transplantation of myogenic stem cells transduced with a therapeutic expression cassette. Development of this method has been hampered by a low frequency of cellular engraftment, the difficulty of tracing transplanted cells, the rapid loss of autologous cells carrying marker genes that are unable to halt muscle necrosis and the difficulty of stable transfer of a large dystrophin gene into myogenic stem cells. We engineered a 5.7 kb miniDys–GFP fusion gene by replacing the dystrophin C-terminal domain ({Delta}CT) with an eGFP coding sequence and removing much of the dystrophin central rod domain ({Delta}H2-R19). In a transgenic mdx4Cv mouse expressing the miniDys–GFP fusion protein under the control of a skeletal muscle-specific promoter, the green fusion protein localized on the sarcolemma, where it assembled the dystrophin–glycoprotein complex and completely prevented the development of dystrophy in transgenic mdx4Cv muscles. When myogenic and other stem cells from these mice were transplanted into mdx4Cv recipients, donor cells can be readily identified in skeletal muscle by direct green fluorescence or by using antibodies against GFP or dystrophin. In mdx4Cv mice reconstituted with bone marrow cells from the transgenic mice, we monitored engraftment in various muscle groups and found the number of miniDys–GFP+ fibers increased with time. We suggest that these transgenic mdx4Cv mice are highly useful for developing autologous cell therapies for DMD.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Duchenne muscular dystrophy (DMD) is characterized primarily by progressive weakness and wasting of muscles and is among the most common genetic disorders. Patients typically are wheelchair-dependent by age 12 and usually die by their early to mid-twenties of respiratory or cardiac failure (1Go). The dystrophin gene, responsible for DMD, is the largest known gene. In muscle, the dystrophin protein nucleates assembly of the dystrophin–glycoprotein complex (DGC), thereby linking the actin cytoskeleton to the extracellular matrix (ECM). Dystrophin and the DGC are thought to protect muscle fibers from mechanical injury by reinforcing the sarcolemmal membrane and/or by dissipating the forces of muscle contraction into the ECM (2Go–5Go). The DGC also has signaling functions, but how these signaling functions protect myofibers from necrosis is poorly understood (6Go). In the absence of dystrophin, the DGC fails to assemble except in very low amounts (7Go).

Two promising approaches for treating DMD are gene and stem cell therapies that could transfer new dystrophin genes to muscles (8Go–11Go). Both strategies have been hampered by the large size of both the dystrophin gene (2.4 Mb) and cDNA (14 kb). Transplantation of myogenic stem cells from a non-autologous donor is a risky procedure that could lead to serious immunological complications. Consequently, ex vivo gene therapy involving transduction of autologous cells with a dystrophin expression vector has emerged as an attractive strategy (12Go). Currently, the best vectors for achieving stable integration of transgenes in stem cells are retroviral vectors, including lentiviruses (12Go–15Go). Lentiviral vectors have a ~8–9 kb cloning capacity, precluding their use for delivering a full-length dystrophin cDNA. Similar vector carrying capacity problems have hampered the development of gene replacement therapy for DMD and have led to the design of a variety of mini- and micro-dystrophin cDNAs (16Go–18Go). Micro-dystrophins <4 kb in size and carrying only four or five of the 24 spectrin-like repeats from the dystrophin central rod domain can be delivered by AAV vectors into muscle, but they are not 100% functional (17Go,19Go–21Go). The smallest dystrophin mini-gene shown to be fully functional ({Delta}H2-R19) (Fig. 1A) has eight spectrin-like repeats, but at 6.8 kb is too large to be stably carried by lentiviral vectors when combined with strong, muscle-specific gene regulatory elements (17Go). We therefore sought to develop improved mini-dystrophin cassettes that would be useful for studies involving stem cell transplantation into dystrophic muscles. In addition to the non-deleterious deletions in the central rod domain, dystrophins lacking the C-terminal (CT) domain have also been shown to be fully functional (22Go). The CT domain of dystrophin interacts with the dystrobrevins and syntrophins, peripheral membrane proteins that form a subcomplex in the DGC (2Go). However, this domain is not essential for the function of dystrophin due to redundant binding sites in the DGC for dystrobrevin and syntrophin (22Go–27Go). We sought to determine whether mini-dystrophins lacking the CT domain would be more functional than the previously described micro-dystrophins (17Go).


Figure 0821
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  		Figure 1. Generation of miniDys–GFP/mdx4Cv transgenic mice. (A) Design and schematic structure of the miniDys–GFP fusion gene. This fusion gene was regulated by the HSA promoter containing a hybrid intron (22Go). A primer pair represented by the two arrows was used to amplify a 470 bp fragment of the fusion-gene transcript. ABD1 and ABD2 (gray shading), N-terminal and central rod actin-binding domains, spectrin-like repeats (white boxes). The basic charged repeats 3, 7, 11–13, 15 and 17 are shaded in gray (repeats 11–17 encompass ABD2); dystrophin hinge domains are indicated by black boxes; DgBD, dystroglycan-binding domain; CR, cysteine-rich domain; E1, exon 1 from the HSA gene; I1/2, 5' portion of the HSA intron 1; Vp1/2, 3' portion of the SV40 VP1 intron. (B) Western analysis of microsome preparations from mdx4Cv, WT and miniDys–GFP/mdx4Cv transgenic mice using an N-terminal dystrophin antibody. (C) A skeletal myofiber bundle from the miniDys–GFP/mdx4Cv transgenic mice displays green fluorescence when directly visualized with a fluorescence microscope. Note that the individual fibers display only weak fluorescence. (D) Cryosections of the quadriceps and diagram muscles from two transgenic mouse lines show uniform expression of the miniDys–GFP fusion protein on the sacrolemma. Scale bar: 100 µm.

 
An ideal therapy for DMD will require systemic delivery of a dystrophin gene, or cells expressing such a gene, to the muscles of the whole body. Accumulating data have suggested that adult stem cells isolated from various types of tissues, such as skeletal muscle, skin, synovial membrane and bone marrow (BM) can home to muscle via the vascular system and engraft in muscle, suggesting that a stem cell therapy could be developed for DMD (12Go,28Go–32Go). Stem cell therapy possesses unique theoretical advantages for treating DMD. First, in the late stages of dystrophy, most myofibers are replaced by fibrous and adipose tissue; therefore, replenishment of myogenic cells is necessary to generate new muscle fibers. Secondly, potential immune responses elicited by viral proteins in vector-mediated gene delivery could be eliminated by ex vivo genetic modification of stem cells isolated from patients before cell transplantation. Thirdly, myogenic and other stem cells can be genetically modified at high efficiency (12Go–15Go). Nonetheless, significant gaps remain in our knowledge of ways to apply such therapies efficiently, including a poor understanding of the molecular mechanisms underlying stem cells proliferation, homing to target tissues and fusing into myofibers.

The efficiency of any stem cell therapy for DMD needs to be significantly improved in animal models before its clinical application. However, assessing the frequency at which dystrophin-expressing cells engraft into muscle following transplantation is hampered by the presence of dystrophin-positive ‘revertant’ fibers that increase in frequency with age in muscles of both mdx mice and cxmd dogs (33Go–38Go). This phenomenon severely interferes with the appraisal of stem cell transplantation studies because current transplant methods generally produce fewer engrafted myofibers than the pre-existing level of revertant fibers (28Go). An easily traceable and functional dystrophin gene would be useful for developing stem cell therapies for DMD.

Here, we describe a 5.7 kb miniDys–GFP fusion gene in which the CT domain of mini-dystrophin ({Delta}H2-R19) was replaced with a green fluorescent protein (eGFP) gene. We evaluated the function of this fusion protein by generating transgenic mice on the mdx4Cv background. Expression was restricted to skeletal muscle by use of the human {alpha}-skeletal actin (HSA) gene regulatory region (22Go,39Go). We found that the miniDys–GFP fusion protein was highly functional and easily detectable on the sarcolemma of skeletal myofibers. When the miniDys–GFP/mdx4Cv transgenic mice were used as donors for primary myoblast, muscle multipotent adult progenitor cell (MAPC) and BM transplantation studies, the green miniDys–GFP+ myofibers were easily detected in skeletal muscles of mdx4Cv recipients. Furthermore, in mdx4Cv BM transplants, we found that the number of miniDys–GFP+ fibers increased in a limited manner with time. These data suggest that this novel miniDys–GFP transgene and the transgenic mice may be useful for experiments aimed at developing cell-based therapies for DMD.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Design of mini-dystrophin/GFP fusion gene
The miniDys–GFP fusion gene was generated by replacing the CT domain of the previously described 6.8 kb mini-dystrophin ({Delta}H2-R19) with the eGFP gene-coding sequence (Fig. 1A). We further decreased the size of this cDNA by removing most of the 5'- and 3'-untranslated regions and also introduced a more optimal Kozak consensus sequence (40Go). The size of final miniDys–GFP gene cDNA was 5.7 kb (see Materials and Methods).

Generation of miniDys–GFP transgenic mice
To assess the functional capacity of the miniDys–GFP fusion protein, we generated an expression vector using the HSA promoter (22Go). Transgenic mice carrying this miniDys–GFP fusion gene were generated and then backcrossed onto the mdx4Cv background. We detected a high level of miniDys–GFP fusion protein by western analysis in skeletal muscles of the miniDys–GFP/mdx4Cv mice (Fig. 1B). Using fluorescence microscopy, green fluorescence could be observed in skeletal muscles of living, neonatal transgenic mice. Dissected skeletal myofiber bundles from the transgenic mice showed strong green fluorescence, but isolated individual myofibers displayed relatively weak fluorescence (Fig. 1C).

Expression pattern and localization of the miniDys–GFP fusion protein
We examined the expression pattern of miniDys–GFP gene in multiple skeletal muscles, heart, liver, spleen, kidney, lung, small intestine, stomach and brain from the transgenic mdx4Cv mice by fluorescence microscopy. Green fluorescence was only observed in skeletal muscles, consistent with the reported expression pattern of the HSA promoter (39Go). Furthermore, we did not detect any miniDys–GFP transcripts by RT–PCR in BM cells isolated from the transgenic mice (discussed subsequently).

Cryosections from the tibialis anterior (TA), extensor digitorum longus (EDL), quadriceps, diaphragm (DPM), heart and liver of transgenic mdx4Cv mice were examined by fluorescence microscopy. All skeletal muscle sections showed uniform expression of the fusion protein on the sarcolemma. Polyclonal antibodies recognizing either the N-terminal domain of dystrophin (41Go) or the GFP displayed perfect co-localization (Fig. 1D) (data not shown). No immunoreactive GFP-fusion protein was observed in any sections from heart or liver (data not shown), again confirming the tissue-specific activity of the HSA promoter.

Morphology of skeletal muscles of miniDys–GFP/mdx4Cv transgenic mice
Hind limb and DPM muscle sections from 2, 6 and 16-month-old miniDys–GFP/mdx4Cv transgenic mice were examined for dystrophic pathology. These muscles displayed normal morphology without any evidence of dystrophy, such as fibrosis, necrosis or mononuclear cell infiltration (Fig. 2A). Control mdx4Cv skeletal muscles displayed a large number of centrally nucleated myofibers resulting from active degeneration and regeneration (42Go,43Go). The number of centrally nucleated myofibers in quadriceps, or DPM, of 2, 6 and 16-month-old miniDys–GFP/mdx4Cv transgenic mice was not different from that of age-matched wild-type (WT) mice (Table 1).


Figure 0822
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  		Figure 2. Morphology of skeletal muscles of miniDys–GFP/mdx4Cv transgenic mice. (A) Hematoxylin and eosin stained quadriceps of 6-month-old miniDys–GFP/mdx4Cv transgenic mice showed no dystrophic features, such as necrotic myofibers, fibrosis or mononuclear cell infiltration as were seen in age-matched mdx4Cv mice. (B and C) Evans blue dye is excluded from DPM (B) and hind limb myofibers (C) in miniDys–GFP/mdx4Cv transgenic mice.

 

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  		Table 1. Percentages of centrally nucleated fibers in mouse skeletal muscles
 
Myofiber integrity and DGC restoration
Damaged and necrotic myofibers in mdx skeletal muscles are permeable to vital dyes such as Evans blue, reflecting a loss of sarcolemmal integrity (44Go). Skeletal myofibers from 6-month-old miniDys–GFP/mdx4Cv transgenic mice, and from age-matched wild type mice (C57BL/6), were not permeable to Evans blue dye. Age-matched mdx4Cv muscles displayed numerous Evans blue dye permeable myofibers (Fig. 2B and C).

All components of the DGC in the mdx skeletal muscles are reduced dramatically (7Go). We tested the hypothesis that expression of the miniDys–GFP fusion protein in mdx4Cv muscles could restore components of DGC onto the sarcolemma. Microsome preparations from skeletal muscles of C57BL/6, mdx4Cv and miniDys–GFP/mdx4Cv transgenic mice were examined by western blotting. The {alpha}-, ß- or {gamma}-sarcoglycan, {alpha}-dystroglycan and {alpha}1-syntrophin were each restored to WT levels in the transgenic mice (Fig. 3A). These DGC components also co-localized with the miniDys–GFP fusion protein on the sarcolemma of myofibers when analyzed by immunostaining (Fig. 3B). The only component of the DGC not restored to normal was nNOS. The nNOS is not properly expressed in any mini- or micro-dystrophin expressing transgenic mice that we have generated, including those expressing the fully functional {Delta}H2-R19 construct (45Go,46Go).


Figure 0823
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  		Figure 3. Expression of the DGC in the skeletal muscles of miniDys–GFP/mdx4Cv transgenic mice. Western blotting analysis of microsome preparation of skeletal muscles (A) and immunostaining of cryosections of quadriceps muscles (B) from WT, mdx4Cv and miniDys–GFP/mdx4Cv transgenic mice using antibodies against ß-dystroglycan, {alpha}-, ß-, {gamma}-sarcoglycan and {alpha}1-syntrophin. Scale bar: 100 µm.

 
Contractile properties of transgenic muscles
No differences were observed in the body masses of the mdx4Cv (35±4 g), transgenic/mdx4Cv (31±5 g) and C57BL/6 (31±2 g) mice, but the EDL and soleus muscle masses of the mdx4Cv mice were 58 and 32% greater, respectively, than those of either WT or transgenic mice (Fig. 4A). The force generating capacity (Po) of EDL muscles from the transgenic mice was 27% lower than that of the mdx4Cv mice and 17% lower than that of the WT mice. The specific Po of the EDL muscles from transgenic mice was intermediate between those of the mdx4Cv and WT mice, which was the only abnormality of note observed for the functional measurements of the transgenic animals. In addition, the specific forces of the soleus and DPM muscles of the transgenic mice were not different from WT and were increased compared with muscles from the mdx4Cv mice (Fig. 4C). The specific Po of DPM muscle strips from the mdx4Cv mice was only 50% of the values of 218 kN/m2 and 227 kN/m2 obtained for the DPM muscles of the transgenic and WT mice (Fig. 4C). Following two 30% lengthening contractions of the EDL and DPM muscles of the mdx4Cv mice, force deficits of 75 and 40%, respectively, were observed. These values were much greater than the force deficits of 10 and 20%, respectively, for the transgenic and 10 and 14%, respectively, for the WT muscles. No differences were observed for the force deficits of the soleus muscles of the mdx4Cv, transgenic and WT mice (Fig. 4D).


Figure 0824
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  		Figure 4. Characteristics of mdx4Cv, miniDys–eGFP/mdx4Cv and WT mice. Muscle masses (A), maximum isometric force (B), specific force (C) and force deficit (D) of three groups of skeletal muscles from mdx4Cv (white bars), miniDys–GFP/mdx4Cv (black bars) and WT (gray bars) mice were compared. SOL, soleus. Data are presented as mean values±standard error from seven to 16 samples. For statistical analysis, an one-way ANOVA (P<0.05) was used to detect differences for each kind of muscle. If a difference was found, additional Student's t-test was performed with a Bonferroni correction. Asterisk indicates a difference from WT; double asterisk indicates a difference from transgenic mice.

 
Transplantation of cells from miniDys–GFP/mdx4Cv transgenic mice into mdx4Cv mice
We investigated whether the use of miniDys–GFP/mdx4Cv transgenic mice as donors for cell transplantation studies would facilitate screening donor-derived dystrophin-positive myofibers in mdx4Cv recipients. We performed intramuscular injections of primary myoblasts or muscle MAPCs, as well as whole BM transplantation to address this question. To reduce host versus graft immune rejection in our transplantation studies, we selected transgenic mice with both major histocompatibility (MHC) loci inherited from the C57BL/6 background.

For myoblast transfer, 5x105 primary myoblasts isolated from the transgenic/mdx4Cv mice were injected into both TA muscles of three mdx4Cv mice. Two weeks later, the muscles were excised and examined for expression of miniDys–GFP. An average of 536±317 (n=6) miniDys–GFP-expressing myofibers were found in each TA muscle. Figure 5A shows numerous miniDys–GFP-expressing myofibers in transplanted, but not in mock-injected TA muscles as determined by fluorescence microscopy. Immunohistochemical staining for dystrophin confirmed the miniDys–GFP expression in recipient myofibers.


Figure 0825
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  		Figure 5. Cell transplantation using miniDys–GFP/mdx4Cv transgenic mouse donors. Primary myoblasts (A) or MAPCs (B) isolated from the skeletal muscles of transgenic mice were injected into mdx4Cv TA muscles. Injected muscles were harvested at 2 weeks. Muscle sections were examined by direct fluorescence and immunostaining against GFP or dystrophin. (A) Scale bar: 500 µm; (B) scale bar for left two pictures: 500 µm; right two pictures: 100 µm. (C) RT–PCR analysis of miniDys–GFP spliced mRNA in various tissues from the mdx4Cv mice transplanted with whole BM cells from transgenic mice 2 months post-transplantation. About 12.5 ng total RNA from the control transgenic TA muscle was used as a positive control. For the other samples, 250 ng total RNA was used. About 50 ng total RNA from each sample was used to amplify ß-actin transcripts as a loading control; see Figure 1A for RT–PCR primer design. Quad., quadriceps; Tg, miniDys–GFP/mdx4Cv mice. (D) Representative miniDys–GFP+ fibers in skeletal muscle sections of BM transplant recipients. Direct fluorescence or immunostaining against GFP or dystrophin was visualized by fluorescence microscopy. Scale bar: 100 µm. (E) The number of miniDys–GFP+ myofibers in the quadriceps of BM transplant recipients increased with time. Transplanted mdx4Cv mice were sacrificed at 2, 4 and 6 months and cryosections were immunostained for GFP. For each time point, one quadriceps muscle from three recipients was examined. For each sample, three cryosections evenly distributed within the muscle were screened and the total numbers of miniDys–GFP+ fibers from those sections were counted.

 
We and others have demonstrated that MAPCs can be isolated from mouse skeletal muscle and can be potentially used as a myogenic cell source (13Go,47Go). We isolated MAPCs from skeletal muscles of the miniDys–GFP/mdx4Cv transgenic mice and injected 7.5x104 MAPCs into mdx4Cv TA muscles. Two weeks later, we harvested injected muscles to screen for miniDys–GFP-expressing myofibers. Ten to 15 GFP+ myofibers were scattered in every muscle section of the transplants. Figure 5B shows a representative region with two miniDys–GFP-expressing fibers. These fibers displayed green fluorescence on the sarcolemma and were immunostained positively for GFP and dystrophin, indicating that donor MAPCs had fused into recipient myofibers.

BM cells have been used for systemic delivery of the dystrophin gene into the skeletal muscles of mdx mice (28Go,48Go,49Go). We lethally irradiated 2-month-old mdx4Cv mice and immediately transplanted whole BM cells isolated from the miniDys–GFP/mdx4Cv transgenic mice. At 2, 4 and 6 months after transplantation, we harvested multiple skeletal muscles, heart, liver and peripheral blood cell from each recipient, and the expression of miniDys–GFP fusion gene in these samples was examined by RT–PCR and direct- or immunofluorescence microscopy.

Harvested tissue extracts were initially analyzed by RT–PCR using primers that specifically amplify a 470 bp miniDys–GFP mRNA fragment that spans an intron in the transgene (Fig. 1A). MiniDys–GFP transcripts were detected in all analyzed skeletal muscles, including the TA, quadriceps, DPM and paraspinal (PS) muscle, but not in heart, liver or peripheral blood from a transplanted mouse sacrificed 2 months after transplantation (Fig. 5C). Note that the DPM muscle displayed relatively low levels of the skeletal {alpha}-actin promoter-driven transgene mRNA. This expression pattern was consistent in tissues from the mice analyzed at 4 and 6 months after transplantation (data not shown).

We next analyzed muscle sections of the BM transplanted mice for miniDys–GFP-positive myofibers. Figure 5D shows representative myofibers expressing the green miniDys–GFP fusion protein. Some myofibers that displayed moderate to weak dystrophin–eGFP expression by immunostaining were not obviously fluorescent when examined by direct fluorescence microscopy (data not shown), suggesting that moderate levels of miniDys–GFP expression are necessary to be detected by direct fluorescence microscopy. We were able to detect miniDys–GFP-positive fibers in cryosections of TA, quadriceps or PS muscles at a frequency of <0.1% of the total myofibers. We did not observe any miniDys–GFP-positive fibers in DPMs from transplanted animals analyzed 2 or 4 months after transplantation. At 6 months post-transplantation, up to two miniDys–GFP+ myofibers were detected in cryosections of the DPMs in three different recipients. This observation was consistent with the relatively low levels of miniDys–GFP transcripts in the DPMs of the mdx transplant recipients (Fig. 5C). No miniDys–GFP+ myofibers were detected in the heart or liver of any recipients (data not shown).

Muscle damage can greatly facilitate incorporation of BM cells into myofibers (31Go,48Go,50Go). The skeletal muscles of mdx mice undergo continuous cycles of myofiber degeneration and regeneration (51Go,52Go). When mdx skeletal muscles are subjected to X-irradiation (as is done prior to BM transplantation), progressive myofiber loss accompanied by increasing fibrosis has been observed (53Go). At 6 months post-transplantation, all of our lethally irradiated mdx4Cv mouse BM transplant recipients were comparatively thin and weak and their harvested skeletal muscles appeared somewhat more dystrophic compared with control mdx4Cv mice (data not shown). We therefore asked whether more miniDys–GFP-expressing fibers were detected in mdx4Cv recipients sacrificed at increasingly longer time points after transplantation. Figure 5E shows a trend of increasing numbers of miniDys–GFP+ myofibers in quadriceps of recipient animals at greater intervals following BM transplantation, although this was not statistically significant. Unfortunately, we were not able to examine BM recipients at longer time points than 6 months due to a high rate of mouse mortality in recipients. The highest average number of miniDys–GFP+ myofibers per 10 µm cryosection of quadriceps muscles was only 10 at 6 months post-transplantation. Taken together, these data indicated that BM cells incorporated into dystrophic mdx4Cv skeletal myofibers at a low frequency, conferring dystrophin expression, and the number of donor-derived dystrophin-positive fibers increased with time. This relatively low level of engraftment is similar to that reported previously by others using WT BM donor cells (28Go,48Go,49Go).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Stem cell transplantation has emerged as a promising approach to developing a treatment for the muscular dystrophies (54Go,55Go). The primary limitation for this approach is a great need for improved efficiencies of cell transplantation into dystrophic models. Consequently, research has focussed on identifying better cell sources for transplantation, understanding the molecular mechanisms by which transplanted cells can home to and engraft in muscle and developing methods for genetically correcting autologous stem cells for therapy (ex vivo gene therapy). We engineered a miniDys–GFP fusion gene to facilitate these studies by enabling easier tracing of donor cells that overexpress a highly functional dystrophin protein. The miniDys–GFP cDNA is only 5.7 kb, such that it can be carried by lentiviral vectors, which are a promising gene transfer vehicle for genetic modification of mdx progenitor cells (12Go,13Go,56Go). Furthermore, we showed that at least three types of stem cells, myoblasts, MAPCs and BM cells could be isolated from the transgenic mdx mice and give rise to miniDys-positive myofibers when transplanted into mdx4Cv mice.

The advantage of using donor cells that express a functional, traceable dystrophin is that engrafted myofibers will be protected from necrosis and display a selective advantage relative to dystrophin-negative myofibers that express a reporter gene such as lacZ. MiniDys-positive myofibers are easily distinguished from revertant dystrophin-positive myofibers by the eGFP moiety. The success of this strategy requires that the miniDys–GFP fusion protein be highly functional, yet small enough to be carried by lentiviral vectors. The construct we used here had not been previously tested, so it was critical to ensure that the miniDys fusion protein was capable of rescuing the mdx mouse muscle phenotype. Our results show that this mini-dystrophin, which lacks two-thirds of the central rod domain as well as the CT domain, was almost fully functional. The only abnormality observed was a slight reduction in force generation in the transgenic mouse EDL muscle. All other transgenic mouse muscles examined displayed normal force development, and the transgenic EDL displayed normal mass, histology and resistance to contraction-induced injury. The reasons for this force drop are not clear, but do not appear to reflect functional deficiencies in the construct because the other muscles were normal. Although dystrophins carrying either the rod or the CT domain truncations have been tested previously in transgenic mice, the present study is the first to test a dystrophin lacking two separate domains in transgenic mdx mice (17Go,22Go). This study is also the first report of a detailed characterization of the physiological and mechanical properties of the 4Cv strain of mdx mice. The mdx4Cv model for DMD has several potential advantages for studying stem cell and gene therapies. First, it displays fewer revertant fibers than do mdx mouse muscles (37Go). Secondly, the mdx4Cv mutation eliminates expression of several non-muscle isoforms of dystrophin, potentially enabling a cleaner analysis of immune responses against exogenous dystrophin (57Go). The mechanical properties of muscles of mdx4Cv mice (Fig. 4) were quite similar to those observed previously for muscles of mdx mice (3Go,58Go).

A previous study also used eGFP as a marker to trace dystrophin expression (59Go). This group fused the GFP-coding sequence with the N-terminus of dystrophin. Although in vivo DNA transfection showed that GFP–dystrophin fusion protein correctly localized on the sarcolemma of myofibers, its overall function in mdx mice was not evaluated. The construct utilized a full-length dystrophin cDNA, precluding its use in lentiviral vectors. The ability to generate stably expressed and proper localizing GFP fusion constructs at both the N-terminal and CT regions of dystrophin highlights the unusual ability of dystrophin to tolerate multiple structural modifications without adversely affecting its functional capacity. In particular, the ability to replace the CT domain of dystrophin with GFP suggests that a variety of other insertions in this region may also be compatible with nearly full dystrophin function.

In the BM transplantation studies using the miniDys–GFP/mdx4Cv transgenic mice as donors, we observed that the number of donor-derived dystrophin-positive fibers in skeletal muscles of mdx4Cv recipients increased with time (Fig. 5E). This increase likely reflects the continuous cycles of myofiber necrosis and regeneration that characterize mdx muscles, providing a niche for ongoing recruitment of BM cells into dystrophic myofibers. Several groups have reported that muscle injury greatly enhances attraction and engraftment of hematopoietic stem cells in muscles of WT mice (31Go,48Go,50Go). The observations presented here are in agreement with those findings. Surprisingly, we also noticed that the DPM, which is the most dystrophic striated muscle in mdx mice (60Go), displayed a very low level of miniDys–GFP mRNA (Fig. 5C) and no detectable donor-derived myofibers even after immunostaining for GFP at 2 or 4 months post-transplantation. At 6 months post-transplantation, the longest time point in this study, recipient skeletal muscles were extremely dystrophic and the number of miniDys–GFP+ fibers remained low. Similar observations were reported when WT mice were used as donors for whole BM transplantation to mdx4Cv mice (49Go). Thus, although a dystrophic microenvironment can enhance donor cell incorporation into muscle, it is not nearly sufficient to lead to a therapeutically significant level of engraftment.

Brazelton et al. (61Go). found that the panniculus carnosus (PC) muscles of WT mouse recipients had the highest frequency of engrafted myofibers following BM transplantation. We were not able to find any donor-derived dystrophin-positive myofibers after thoroughly screening the PC sections of our mdx4Cv recipients (data not shown). This discrepancy may reflect different microenvironments in the skeletal muscles of mdx4Cv and WT mice or may be due to strain differences.

The mechanisms by which BM stem cells incorporate into myofibers remain unclear. LaBarge and Blau (50Go) reported that donor hematopoietic cells can form myofibers via the canonical myogenic pathway. Other groups have suggested that the low frequency of donor-derived myofibers occurred via fusion of myeloid cells or BM stromal cells with myofibers (48Go,62Go). This latter study did not find any donor-derived myoblast colonies when mononuclear muscle cells were cultured in vitro (48Go), nor we have been able to isolate such colonies from our mdx4Cv BM transplant recipients (data not shown). However, high levels of infiltrating macrophages and neutrophils are found in injured or dystrophic muscles (63Go,64Go) and myeloid cells have been observed to fuse with myofibers in vivo (48Go). Interestingly, IL-4 has been shown to facilitate cell–cell fusion of myoblasts to myotubes as well as macrophage fusion to form giant cells (65Go–67Go). In addition, the fusion of BM cells to hepatocytes, cardiomyocytes and Purkinje neurons after BM transplantation has been reported by several groups (68Go–71Go). Further studies to understand the underlying mechanisms of myofiber engraftment are clearly needed to improve the efficiency of stem cell strategies for DMD, and such studies may be aided by the reagents described in this paper.

In summary, the miniDys-GFP gene encodes a small and easily traceable, but highly functional protein. Transgenic/mdx4Cv mice overexpress this fusion protein exclusively in skeletal muscles, providing a rich source of cells with which to develop and optimize cell transplant therapies for DMD. Transplanted cells can be readily distinguished from revertant mdx myofibers, and the expressed dystrophin provides functional rescue and selection for engrafted myofibers. The same miniDys–GFP expression cassette is small enough to be readily cloned into lentiviral vectors, making it a potentially useful construct with which to study approaches for ex vivo gene therapy of DMD using autologous cell transplants. A considerable increase in our understanding of the best cell source for transplants, optimal methods to isolate or propagate those cells and the mechanisms underlying cell homing and engraftment in muscle are needed before cell therapies can be useful for the muscular dystrophies. The miniDys–GFP gene and transgenic mice may help facilitate these studies.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
miniDys–GFP/mdx4Cv transgenic mice
A miniDys–GFP fusion gene was generated in several cloning steps. We first replaced TAGGAA (underlined nucleotides are the stop codon of dystrophin gene) with TCTAGA (XbaI site) in pSV40pA{Delta}71-78 (17Go) using the QuikChange® XL Site-Directed Mutagenesis Kit (Stratagene) (forward primer: 5'-GGAAACTGACACAATTCTAGAAGTCTTTTCCAC-3'; reverse primer: 5'-GTGGAAAAGACTTCTAGAATTGTGTCAGTTTCC-3'). A PCR-amplified XbaI-flanked eGFP-coding sequence without its translational initiation region (forward primer: 5'-GCTCTAGAGGTGAGCAAGGGCGAGGAG-3'; reverse primer: 5'-CCTCTAGAATTCCGGCCGCTTT AC-3') was then inserted in frame to generate pSV40pA{Delta}71-78-GFP. A 2222-bp HindIII fragment from this plasmid was used to replace the 1478-bp HindIII fragment in the pCK6-{Delta}R4-R23{Delta}71-78 (17Go) to generate pCK6-{Delta}R4-R23{Delta}71-78-GFP. We then replaced the 2546-bp NsiI–BspEI fragment in the pCK6-{Delta}R4-R23{Delta}71-78-GFP with a 3929 bp NsiI–BspEI fragment from pBSX-{Delta}H2-R19 (17Go) to generate pCK6{Delta}H2-R19{Delta}71-78-eGFP (CK6-miniDys–eGFP). Finally, a 5128-bp EagI fragment containing {Delta}H2-R19{Delta}71-78-eGFP (miniDys–eGFP) in pCK6{Delta}H2-R19{Delta}71-78-GFP was inserted into pHSAvpSV40p expression vector (22Go) at the NotI site to generate pHSA-{Delta}H2-R19{Delta}71-78-GFP-pA. The excised miniDys–GFP expression cassette was injected into B6C3F1 hybrid fertilized eggs (Taconic), and F0 mice were screened for the transgene by PCR. Four positive F0 mice were backcrossed onto the B6Ros.Cg-Dmdmdx-4Cv (‘mdx4Cv’) background. Most studies focussed on the line 18916 that had the most uniform transgene expression levels in skeletal muscles.

Morphological assays
Quadriceps, TA, EDL and DPM muscles were removed from mice, embedded in O.C.T. medium (Tissue-Tek), frozen in liquid nitrogen and cut into 5 µm sections. After fixation in 3.7% formaldehyde, sections were stained in hematoxylin and eosin-phloxine. Stained sections were imaged with a Nikon E1000 microscope connected to a Spot-2 CCD camera. To determine the percentage of myofibers containing central nuclei, the number of muscle fibers with centrally located nuclei was divided by the total number of muscle fibers. Evans blue assays were as described previously (72Go).

Immunofluorescence analysis
As described earlier, 5 µm cryosections of various tissues were prepared. Immunofluorescence detection was performed with primary antibodies recognizing the following proteins: the N-terminal domain of dystrophin (41Go), green fluorescence protein (Molecular Probes), ß-dystroglycan and {alpha}-, ß-, {gamma}-sarcoglycan (Novacastra Laboratories, Ltd) and {alpha}1-syntrophin (73Go). After incubation with primary antibodies, the cryosections were incubated with a goat anti-rabbit or goat anti-mouse secondary antibody conjugated with Alexa 594 (Molecular Probes). Images were collected on a Nikon E1000 microscope under identical conditions using a Spot-2 CCD camera.

Measurement of contractile properties
Contractile properties were measured in vitro on EDL and soleus muscles and DPM strips obtained from 7–10-month-old WT C57BL/6, mdx4Cv and miniDys–GFP/mdx4Cv transgenic male mice by methods described previously (74Go). An exception was the control data on DPM strips that were obtained from 17–18-month-old female WT mice. Control data on DPM strips of WT mice do not differ among mice from 6 to 24 months of age and between males and females (75Go). EDL and soleus muscles were evaluated from only one leg. In all, seven EDL, seven soleus and 16 DPM muscles were examined from the mdx4Cv mice, seven EDL, eight soleus and 16 DPM muscles from the transgenic mice and eight EDL, eight soleus and 15 DPM from WT mice. Mice were anesthetized with an intraperitoneal injection of 1.3% avertin (0.015 ml/g body mass) with supplemental injections as required to prevent response to tactile stimuli. EDL and soleus muscles were isolated and proximal and distal tendons were tied firmly with silk (5.0) suture. Entire DPM muscles were excised. Muscles were removed from the mouse and submerged in a bath that contained buffered mammalian Ringer's solution (75Go). The intact fiber lengths of DPM strips extended from the central tendon to the rib. Ties were placed firmly around both the central tendon and the rib. For each of the three muscles, with the muscle in a bath and platinum electrodes placed on either side, one end of the tendon was tied to a force transducer (model BG-50, Kulite Semiconductor Products) and the other tendon to the lever arm of a servomotor (model 305B Aurora Scientific, Richmond Hill, ON, Canada). Muscles were stimulated directly with a pulse duration of 2 ms. With the muscle at resting length, the voltage was increased to produce a maximum twitch force and muscle length was adjusted to optimum length (Lo) for force development. With the muscle length set at Lo, the stimulation frequency was increased until the development of force plateaued at the maximum isometric tetanic force (Po). Stimulation durations were of 300 ms for EDL muscles and 900 ms for soleus and DPM muscles. Po was usually occurred at a frequency of ~180 Hz for EDL and ~150 Hz for soleus and DPM muscles. The susceptibility of muscles to contraction-induced injury was assessed by two lengthening contractions. The muscles were set at Lo, activated maximally, and then stretched through a strain of 30% at velocity of 1 Lf/s and then returned at the same velocity to Lo, relaxed for a 10 s recovery period and then exposed to a second stretch of 30%. The muscles were then allowed to recover for 1 min before the maximum force was measured. Muscles were removed from the bath, trimmed, blotted, weighed and subsequently frozen in isopentane cooled by dry ice. The total fiber cross-sectional area (CSA, cm2) was calculated based on the measurements of optimal muscle length (mm), muscle mass (mg), a muscle density of 1.06 g/cm2 and an Lf/Lo ratio of 0.44 for the EDL, 0.71 for the soleus and 1 for the DPM muscles (74Go). The specific Po (kN/m2) was determined by dividing Po (kN) by CSA (m2). The force deficit produced by the lengthening contraction protocol was assessed by expressing the Po (mN) measured after the two lengthening contraction protocol as a percentage of the Po (mN) before injury.

Western analysis
Skeletal muscle microsomes from 4-month-old C57BL/6, mdx4Cv and miniDys–GFP/mdx4Cv transgenic mice (line 18916) were prepared as described previously (76Go) and used for western analysis. The final microsome pellet was resuspended in 0.3 M sucrose and 20 mM Tris-maleate (pH 7.0). Protein concentrations of each sample were measured by the Bradford assay (77Go), and equal protein loading was verified by SDS–PAGE. KCl-washed microsomes were electrophoretically separated on 4–12% gradient SDS–PAGE polyacrylamide gels (BioRad) and analyzed by western blot using the primary antibodies used for immunostaining, described earlier.

Detection of MHC loci
The MHC loci in C57BL/6 and C3 mouse strains can be distinguished by PCR amplification of two markers (D17Mit10 and D17Mit51) around the MHC locus (http://jaxmice.jax.org). F2 mice from line 18916 were examined by PCR and males carrying the two C57BL/6 markers were chosen for further backcrossing with female B6Ros.Cg-Dmdmdx-4Cv mice (‘mdx4Cv’) (Jackson Laboratory, Bar Harbor, ME, USA) for seven generations prior to using them for cell transplantation into mdx4Cv mouse recipients.

Preparation of primary myoblasts and intramuscular transplantation
All limb muscles from 1-month-old F5 miniDys–GFP/mdx4Cv transgenic mice were excised, minced and digested with 0.2% Collagenase II and 1.2 U/ml Dispase in 1xphosphate-buffered saline (PBS) (pH 7.2) with 1 mM CaCl2 at 37°C for 45 min. The digestion was stopped by adding F10 (GIBCO BRL) with 15% horse serum (Atlanta Biologicals). Digested muscle was filtered through 70 µm and then 40 µm nylon filters (BD Falcon). Mononuclear cells were cultured in gelatin-coated plates (0.67%) with F10C (GIBCO BRL) supplemented with 15% horse serum (Atlanta Biologicals) and 5 ng/ml basic FGF2 (R & D system) for 48 h to enrich for myoblasts. Cells were trypsinized, washed once and resuspended in 1xPBS (pH 7.2). For transplantation, 2-month-old mdx4Cv mice were anesthetized via sustained inhalation of isoflurane in medical oxygen at a concentration sufficient to eliminate response to tactile stimuli. Their TA muscles were surgically exposed. Forty microliters containing 5x105 cells suspension was loaded in 1 cc syringes and injected longitudinally into each TA muscle. After injection, the skin incision was closed with cyanoacrylate adhesive (Nexaband S/C, World Precision Instruments, Sarasota, FL, USA).

Isolation of muscle MAPCs
Limb muscles of miniDys–GFP/mdx4Cv transgenic mice were excised and mononuclear muscle cells were prepared as described above. We immediately cultured these cells in mouse MAPC media as described to obtain muscle MAPCs (47Go). Before intramuscular injection, the multipotent differentiation capacity of isolated MAPCs was examined by in vitro differentiation assays. The MAPCs were trypsinized, washed and resuspended in 1xPBS (pH 7.2). As described above, 7.5x104 MAPCs were intramuscularly injected into each TA muscle of 2-month-old mdx4Cv mice.

Intravenous transplantation of whole BM cells
Whole BM cells were sterilely collected by a slightly modified method described by LaBarge and Blau (50Go). Briefly, 6–8-week-old F5 miniDys–GFP/mdx4Cv transgenic mice were sacrificed by cervical dislocation, briefly immersed in 75% ethanol and had their skin peeled back from a midline, circumferential incision. The femurs were dissociated and the muscles around them removed, and the marrow cavities were flushed with a 26-gage needle containing ice-cold Iscove's DMD medium (GIBCO BRL) with 2% fetal bovine serum (FBS). Marrow cells were further dissociated by triturating through the 26-gage needles. The resulting single-cell suspension was spun at 300g for 10 min, and the pellet was resuspended in ice-cold Iscove's DMD with 2% FBS to 10 million nucleated cells per ml. Two-month-old female mdx4Cv mice were then lethally irradiated (1100 rad) and immediately injected with 200 µl of 2 million nucleated cells via tail vein.

RT–PCR amplification of miniDys–eGFP mRNA
Total RNA from TA, quadriceps (Quad), DPM, PS muscle, heart, liver or peripheral blood of recipients were isolated with an RNease Mini Kit (Qiagen), respectively, according to manufacturer's instructions. Primer pairs (forward primer: 5'-AGCCGAGAGTAGCAGTTGTAGC-3'; reverse primer: 5'-TTACATTTTTGACCTGCCAGTG-3') were designed (Fig. 1A) to specifically amplify a 470 bp fragment from the miniDys–GFP mRNA, and another primer pair (forward primer: 5'-TGTGACGTTGACATCCGTAAAG-3'; reverse primer: 5'-AAACGCAGCTCAGTAACAGTCC) was used to amplify a 300 bp fragment of ß-actin mRNA. QIAGEN OneStep RT–PCR kits were used to detect both miniDys–eGFP and ß-actin mRNAs. For amplifying ß-actin mRNA fragments (loading control), 25 ng total RNA from each sample was used, whereas for miniDys–eGFP transcripts, 250 ng total RNA was used.


   ACKNOWLEDGEMENTS
 
We thank X. Ye and G. Stamatoyannopoulos for the generation of MiniDys–GFP transgenic mice and J. Yan, M. Weinreich, J.C. Angello, C.A. Blau and S.D. Hauschka for advice and technical assistance. We also thank all members in the Chamberlain lab for helpful discussions and advice. These studies were supported by grants from the National Institutes of Health (P01 AG015434, P01 NS46788 and P01 AS046788) and the Muscular Dystrophy Association (USA) (to J.S.C. and J.A.F.). S.L. was supported by a Research Development Grant from the Muscular Dystrophy Association (USA).

Conflict of Interest statement. None declared.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

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