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

A highly functional mini-dystrophin/GFP fusiongene for cell and gene therapy studies ofDuchenne 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

Received February 16, 2006; Revised and Accepted March 25, 2006

A promising approach for treating Duchenne muscular dystrophy (DMD) is by autologous cell transplantationof myogenic stem cells transduced with a therapeutic expression cassette. Development of this method hasbeen hampered by a low frequency of cellular engraftment, the difficulty of tracing transplanted cells, therapid loss of autologous cells carrying marker genes that are unable to halt muscle necrosis and the difficultyof stable transfer of a large dystrophin gene into myogenic stem cells. We engineered a 5.7 kb miniDys–GFPfusion gene by replacing the dystrophin C-terminal domain (DCT) with an eGFP coding sequence and remov-ing much of the dystrophin central rod domain (DH2-R19). In a transgenic mdx4Cv mouse expressing theminiDys–GFP fusion protein under the control of a skeletal muscle-specific promoter, the green fusionprotein localized on the sarcolemma, where it assembled the dystrophin–glycoprotein complex and comple-tely prevented the development of dystrophy in transgenic mdx4Cv muscles. When myogenic and other stemcells from these mice were transplanted into mdx4Cv recipients, donor cells can be readily identified in skel-etal muscle by direct green fluorescence or by using antibodies against GFP or dystrophin. In mdx4Cv micereconstituted with bone marrow cells from the transgenic mice, we monitored engraftment in various musclegroups and found the number of miniDys–GFP1 fibers increased with time. We suggest that these transgenicmdx4Cv mice are highly useful for developing autologous cell therapies for DMD.

INTRODUCTION

Duchenne muscular dystrophy (DMD) is characterizedprimarily by progressive weakness and wasting of musclesand is among the most common genetic disorders. Patientstypically are wheelchair-dependent by age 12 and usuallydie by their early to mid-twenties of respiratory or cardiacfailure (1). The dystrophin gene, responsible for DMD, isthe largest known gene. In muscle, the dystrophin proteinnucleates assembly of the dystrophin–glycoprotein complex(DGC), thereby linking the actin cytoskeleton to the extra-cellular matrix (ECM). Dystrophin and the DGC are thoughtto protect muscle fibers from mechanical injury by reinforcing

the sarcolemmal membrane and/or by dissipating the forces ofmuscle contraction into the ECM (2–5). The DGC also hassignaling functions, but how these signaling functionsprotect myofibers from necrosis is poorly understood (6). Inthe absence of dystrophin, the DGC fails to assemble exceptin very low amounts (7).

Two promising approaches for treating DMD are gene andstem cell therapies that could transfer new dystrophin genesto muscles (8–11). Both strategies have been hampered bythe large size of both the dystrophin gene (2.4 Mb) andcDNA (14 kb). Transplantation of myogenic stem cells froma non-autologous donor is a risky procedure that could leadto serious immunological complications. Consequently,

# The Author 2006. Published by Oxford University Press. All rights reserved.For Permissions, please email: [email protected]

*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: [email protected]

Human Molecular Genetics, 2006, Vol. 15, No. 10 1610–1622doi:10.1093/hmg/ddl082Advance Access published on April 4, 2006

by guest on September 27, 2013

http://hmg.oxfordjournals.org/

Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from



Dow

nloaded from














Human Molecular Genetics, 2006, Vol. 15, No. 10 1611

ex vivo gene therapy involving transduction of autologouscells with a dystrophin expression vector has emerged asan attractive strategy (12). Currently, the best vectors forachieving stable integration of transgenes in stem cells areretroviral vectors, including lentiviruses (12–15). Lentiviralvectors have a �8–9 kb cloning capacity, precluding theiruse for delivering a full-length dystrophin cDNA. Similarvector carrying capacity problems have hampered the develop-ment of gene replacement therapy for DMD and have ledto the design of a variety of mini- and micro-dystrophincDNAs (16–18). Micro-dystrophins ,4 kb in size and carry-ing only four or five of the 24 spectrin-like repeats from thedystrophin central rod domain can be delivered by AAVvectors into muscle, but they are not 100% functional(17,19–21). The smallest dystrophin mini-gene shown to befully functional (DH2-R19) (Fig. 1A) has eight spectrin-likerepeats, but at 6.8 kb is too large to be stably carried by lenti-viral vectors when combined with strong, muscle-specificgene regulatory elements (17). We therefore sought todevelop improved mini-dystrophin cassettes that would beuseful for studies involving stem cell transplantation intodystrophic muscles. In addition to the non-deleteriousdeletions in the central rod domain, dystrophins lackingthe C-terminal (CT) domain have also been shown tobe fully functional (22). The CT domain of dystrophininteracts with the dystrobrevins and syntrophins, peripheralmembrane proteins that form a subcomplex in the DGC(2). However, this domain is not essential for the functionof dystrophin due to redundant binding sites inthe DGC for dystrobrevin and syntrophin (22–27). Wesought to determine whether mini-dystrophins lacking the CTdomain would be more functional than the previouslydescribed micro-dystrophins (17).

An ideal therapy for DMD will require systemic deliveryof a dystrophin gene, or cells expressing such a gene, tothe muscles of the whole body. Accumulating data havesuggested that adult stem cells isolated from various typesof tissues, such as skeletal muscle, skin, synovial membraneand bone marrow (BM) can home to muscle via the vascularsystem and engraft in muscle, suggesting that a stem celltherapy could be developed for DMD (12,28–32). Stemcell therapy possesses unique theoretical advantages for treat-ing DMD. First, in the late stages of dystrophy, most myo-fibers are replaced by fibrous and adipose tissue; therefore,replenishment of myogenic cells is necessary to generatenew muscle fibers. Secondly, potential immune responses eli-cited by viral proteins in vector-mediated gene delivery couldbe eliminated by ex vivo genetic modification of stem cellsisolated from patients before cell transplantation. Thirdly,myogenic and other stem cells can be genetically modified

at high efficiency (12–15). Nonetheless, significant gapsremain in our knowledge of ways to apply such therapies effi-ciently, including a poor understanding of the molecularmechanisms underlying stem cells proliferation, homing totarget tissues and fusing into myofibers.

The efficiency of any stem cell therapy for DMD needs tobe significantly improved in animal models before its clinicalapplication. However, assessing the frequency at whichdystrophin-expressing cells engraft into muscle followingtransplantation is hampered by the presence of dystrophin-positive ‘revertant’ fibers that increase in frequency with agein muscles of both mdx mice and cxmd dogs (33–38). Thisphenomenon severely interferes with the appraisal of stemcell transplantation studies because current transplantmethods generally produce fewer engrafted myofibers thanthe pre-existing level of revertant fibers (28). An easily trace-able and functional dystrophin gene would be useful fordeveloping stem cell therapies for DMD.

Here, we describe a 5.7 kb miniDys–GFP fusion gene inwhich the CT domain of mini-dystrophin (DH2-R19) wasreplaced with a green fluorescent protein (eGFP) gene. Weevaluated the function of this fusion protein by generatingtransgenic mice on the mdx4Cv background. Expression wasrestricted to skeletal muscle by use of the human a-skeletalactin (HSA) gene regulatory region (22,39). We found thatthe miniDys–GFP fusion protein was highly functional andeasily detectable on the sarcolemma of skeletal myofibers.When the miniDys–GFP/mdx4Cv transgenic mice were usedas donors for primary myoblast, muscle multipotent adult pro-genitor cell (MAPC) and BM transplantation studies, the greenminiDys–GFPþ myofibers were easily detected in skeletalmuscles of mdx4Cv recipients. Furthermore, in mdx4Cv BMtransplants, we found that the number of miniDys–GFPþ

fibers increased in a limited manner with time. These datasuggest that this novel miniDys–GFP transgene and the trans-genic mice may be useful for experiments aimed at developingcell-based therapies for DMD.

RESULTS

Design of mini-dystrophin/GFP fusion gene

The miniDys–GFP fusion gene was generated by replacingthe CT domain of the previously described 6.8 kb mini-dystrophin (DH2-R19) with the eGFP gene-coding sequence(Fig. 1A). We further decreased the size of this cDNA byremoving most of the 50- and 30-untranslated regions andalso introduced a more optimal Kozak consensus sequence(40). The size of final miniDys–GFP gene cDNA was5.7 kb (see Materials and Methods).

Figure 1. Generation of miniDys–GFP/mdx4Cv transgenic mice. (A) Design and schematic structure of the miniDys–GFP fusion gene. This fusion gene wasregulated by the HSA promoter containing a hybrid intron (22). A primer pair represented by the two arrows was used to amplify a 470 bp fragment of thefusion-gene transcript. ABD1 and ABD2 (gray shading), N-terminal and central rod actin-binding domains, spectrin-like repeats (white boxes). The basiccharged 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, 50 portion of the HSA intron 1; Vp1/2, 30 portion ofthe SV40 VP1 intron. (B) Western analysis of microsome preparations from mdx4Cv, WT and miniDys–GFP/mdx4Cv transgenic mice using an N-terminal dys-trophin antibody. (C) A skeletal myofiber bundle from the miniDys–GFP/mdx4Cv transgenic mice displays green fluorescence when directly visualized with afluorescence microscope. Note that the individual fibers display only weak fluorescence. (D) Cryosections of the quadriceps and diagram muscles from twotransgenic mouse lines show uniform expression of the miniDys–GFP fusion protein on the sacrolemma. Scale bar: 100 mm.

1612 Human Molecular Genetics, 2006, Vol. 15, No. 10

Generation of miniDys–GFP transgenic mice

To assess the functional capacity of the miniDys–GFP fusionprotein, we generated an expression vector using the HSApromoter (22). Transgenic mice carrying this miniDys–GFPfusion gene were generated and then backcrossed onto themdx4Cv background. We detected a high level of miniDys–GFP fusion protein by western analysis in skeletal musclesof the miniDys–GFP/mdx4Cv mice (Fig. 1B). Using fluor-escence microscopy, green fluorescence could be observedin skeletal muscles of living, neonatal transgenic mice. Dis-sected skeletal myofiber bundles from the transgenic miceshowed strong green fluorescence, but isolated individualmyofibers displayed relatively weak fluorescence (Fig. 1C).

Expression pattern and localization of theminiDys–GFP fusion protein

We examined the expression pattern of miniDys–GFP gene inmultiple skeletal muscles, heart, liver, spleen, kidney, lung,small intestine, stomach and brain from the transgenicmdx4Cv mice by fluorescence microscopy. Green fluorescencewas only observed in skeletal muscles, consistent with thereported expression pattern of the HSA promoter (39). Fur-thermore, we did not detect any miniDys–GFP transcriptsby RT–PCR in BM cells isolated from the transgenic mice(discussed subsequently).

Cryosections from the tibialis anterior (TA), extensor digi-torum longus (EDL), quadriceps, diaphragm (DPM), heartand liver of transgenic mdx4Cv mice were examined by

fluorescence microscopy. All skeletal muscle sectionsshowed uniform expression of the fusion protein on the sarco-lemma. Polyclonal antibodies recognizing either the N-terminal domain of dystrophin (41) or the GFP displayedperfect co-localization (Fig. 1D) (data not shown). Noimmunoreactive GFP-fusion protein was observed in any sec-tions from heart or liver (data not shown), again confirmingthe 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 and16-month-old miniDys–GFP/mdx4Cv transgenic mice wereexamined for dystrophic pathology. These muscles displayednormal morphology without any evidence of dystrophy, suchas fibrosis, necrosis or mononuclear cell infiltration(Fig. 2A). Control mdx4Cv skeletal muscles displayed a largenumber of centrally nucleated myofibers resulting fromactive degeneration and regeneration (42,43). The number ofcentrally nucleated myofibers in quadriceps, or DPM, of 2, 6and 16-month-old miniDys–GFP/mdx4Cv transgenic micewas not different from that of age-matched wild-type (WT)mice (Table 1).

Myofiber integrity and DGC restoration

Damaged and necrotic myofibers in mdx skeletal muscles arepermeable to vital dyes such as Evans blue, reflecting a loss of

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-matchedmdx4Cv mice. (B and C) Evans blue dye is excluded from DPM (B) and hind limb myofibers (C) in miniDys–GFP/mdx4Cv transgenic mice.


sarcolemmal integrity (44). Skeletal myofibers from6-month-old miniDys–GFP/mdx4Cv transgenic mice, andfrom age-matched wild type mice (C57BL/6), were notpermeable to Evans blue dye. Age-matched mdx4Cv musclesdisplayed numerous Evans blue dye permeable myofibers(Fig. 2B and C).

All components of the DGC in the mdx skeletal muscles arereduced dramatically (7). We tested the hypothesis thatexpression of the miniDys–GFP fusion protein in mdx4Cv

muscles could restore components of DGC onto the sarco-lemma. Microsome preparations from skeletal muscles ofC57BL/6, mdx4Cv and miniDys–GFP/mdx4Cv transgenicmice were examined by western blotting. The a-, b- org-sarcoglycan, a-dystroglycan and a1-syntrophin were each

restored to WT levels in the transgenic mice (Fig. 3A).These DGC components also co-localized with theminiDys–GFP fusion protein on the sarcolemma of myofiberswhen analyzed by immunostaining (Fig. 3B). The only com-ponent of the DGC not restored to normal was nNOS. ThenNOS is not properly expressed in any mini- or micro-dystrophin expressing transgenic mice that we have generated,including those expressing the fully functional DH2-R19construct (45,46).

Contractile properties of transgenic muscles

No differences were observed in the body masses of themdx4Cv (35 + 4 g), transgenic/mdx4Cv (31 + 5 g) and

Figure 3. Expression of the DGC in the skeletal muscles of miniDys–GFP/mdx4Cv transgenic mice. Western blotting analysis of microsome preparation ofskeletal muscles (A) and immunostaining of cryosections of quadriceps muscles (B) from WT, mdx4Cv and miniDys–GFP/mdx4Cv transgenic mice usingantibodies against b-dystroglycan, a-, b-, g-sarcoglycan and a1-syntrophin. Scale bar: 100 mm.

Table 1. Percentages of centrally nucleated fibers in mouse skeletal muscles

Muscle type Two months old Six months old Sixteen months old

WT mdx4Cv MiniDys–GFP WT mdx4Cv MiniDys–GFP WT mdx4Cv MiniDys–GFP

Quad ,1 53 ,1 ,1 82 2 ND 54 ,1DPM ,1 ND ND ,1 62 ,1 ND 39 ,1

WT, C57BL/6; mdx4Cv, transgene negative mice; MiniDys–GFP, transgene positive mdx4Cv mice; Quad, quadriceps; ND, not determined. The mdx4Cv

and MiniDys–GFP mice were progenies from the same breeding mice. The numbers are averages of three sections.


C57BL/6 (31 + 2 g) mice, but the EDL and soleus musclemasses 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 EDLmuscles from the transgenic mice was 27% lower thanthat of the mdx4Cv mice and 17% lower than that of theWT mice. The specific Po of the EDL muscles from trans-genic mice was intermediate between those of the mdx4Cv

and WT mice, which was the only abnormality of noteobserved for the functional measurements of the transgenicanimals. In addition, the specific forces of the soleus andDPM muscles of the transgenic mice were not differentfrom WT and were increased compared with muscles fromthe mdx4Cv mice (Fig. 4C). The specific Po of DPMmuscle strips from the mdx4Cv mice was only 50% of thevalues of 218 kN/m2 and 227 kN/m2 obtained for the DPMmuscles of the transgenic and WT mice (Fig. 4C). Followingtwo 30% lengthening contractions of the EDL and DPMmuscles of the mdx4Cv mice, force deficits of 75 and 40%,respectively, were observed. These values were muchgreater than the force deficits of 10 and 20%, respectively,for the transgenic and 10 and 14%, respectively, forthe WT muscles. No differences were observed for theforce deficits of the soleus muscles of the mdx4Cv, transgenicand WT mice (Fig. 4D).

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 studieswould facilitate screening donor-derived dystrophin-positivemyofibers in mdx4Cv recipients. We performed intramuscularinjections of primary myoblasts or muscle MAPCs, as well aswhole BM transplantation to address this question. To reducehost versus graft immune rejection in our transplantationstudies, we selected transgenic mice with both major histocom-patibility (MHC) loci inherited from the C57BL/6 background.

For myoblast transfer, 5 � 105 primary myoblasts isolatedfrom the transgenic/mdx4Cv mice were injected into both TAmuscles of three mdx4Cv mice. Two weeks later, the muscleswere excised and examined for expression of miniDys–GFP.An average of 536 + 317 (n ¼ 6) miniDys–GFP-expressingmyofibers were found in each TA muscle. Figure 5A showsnumerous miniDys–GFP-expressing myofibers in trans-planted, but not in mock-injected TA muscles as determinedby fluorescence microscopy. Immunohistochemical stainingfor dystrophin confirmed the miniDys–GFP expression inrecipient myofibers.

We and others have demonstrated that MAPCs can be iso-lated from mouse skeletal muscle and can be potentially

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 differ-ences for each kind of muscle. If a difference was found, additional Student’s t-test was performed with a Bonferroni correction. Asterisk indicates a differencefrom WT; double asterisk indicates a difference from transgenic mice.


Figure 5. Cell transplantation using miniDys–GFP/mdx4Cv transgenic mouse donors. Primary myoblasts (A) or MAPCs (B) isolated from the skeletal musclesof transgenic mice were injected into mdx4Cv TA muscles. Injected muscles were harvested at 2 weeks. Muscle sections were examined by direct fluorescenceand immunostaining against GFP or dystrophin. (A) Scale bar: 500 mm; (B) scale bar for left two pictures: 500 mm; right two pictures: 100 mm. (C) RT–PCRanalysis 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 RNAwas used. About 50 ng total RNA from each sample was used to amplify b-actin transcripts as a loading control; see Figure 1A for RT–PCR primerdesign. Quad., quadriceps; Tg, miniDys–GFP/mdx4Cv mice. (D) Representative miniDys–GFPþ 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 mm. (E) The number ofminiDys–GFPþ myofibers in the quadriceps of BM transplant recipients increased with time. Transplanted mdx4Cv mice were sacrificed at 2, 4 and 6months and cryosections were immunostained for GFP. For each time point, one quadriceps muscle from three recipients was examined. For eachsample, three cryosections evenly distributed within the muscle were screened and the total numbers of miniDys–GFPþ fibers from those sections werecounted.


used as a myogenic cell source (13,47). We isolated MAPCsfrom skeletal muscles of the miniDys–GFP/mdx4Cv transgenicmice and injected 7.5 � 104 MAPCs into mdx4Cv TA muscles.Two weeks later, we harvested injected muscles to screen forminiDys–GFP-expressing myofibers. Ten to 15 GFPþ myofi-bers were scattered in every muscle section of the transplants.Figure 5B shows a representative region with two miniDys–GFP-expressing fibers. These fibers displayed greenfluorescence on the sarcolemma and were immunostainedpositively for GFP and dystrophin, indicating that donorMAPCs had fused into recipient myofibers.

BM cells have been used for systemic delivery of thedystrophin gene into the skeletal muscles of mdx mice(28,48,49). We lethally irradiated 2-month-old mdx4Cv

mice and immediately transplanted whole BM cells isolatedfrom the miniDys–GFP/mdx4Cv transgenic mice. At 2, 4 and6 months after transplantation, we harvested multiple skeletalmuscles, heart, liver and peripheral blood cell from each reci-pient, and the expression of miniDys–GFP fusion gene inthese samples was examined by RT–PCR and direct- orimmunofluorescence microscopy.

Harvested tissue extracts were initially analyzed by RT–PCR using primers that specifically amplify a 470 bpminiDys–GFP mRNA fragment that spans an intron in thetransgene (Fig. 1A). MiniDys–GFP transcripts were detectedin all analyzed skeletal muscles, including the TA, quadri-ceps, DPM and paraspinal (PS) muscle, but not in heart,liver or peripheral blood from a transplanted mouse sacri-ficed 2 months after transplantation (Fig. 5C). Note thatthe DPM muscle displayed relatively low levels of the skel-etal a-actin promoter-driven transgene mRNA. Thisexpression pattern was consistent in tissues from the miceanalyzed at 4 and 6 months after transplantation (data notshown).

We next analyzed muscle sections of the BM transplantedmice for miniDys–GFP-positive myofibers. Figure 5Dshows representative myofibers expressing the greenminiDys–GFP fusion protein. Some myofibers that displayedmoderate to weak dystrophin–eGFP expression by immuno-staining were not obviously fluorescent when examined bydirect fluorescence microscopy (data not shown), suggestingthat moderate levels of miniDys–GFP expression are necess-ary to be detected by direct fluorescence microscopy. We wereable to detect miniDys–GFP-positive fibers in cryosections ofTA, quadriceps or PS muscles at a frequency of ,0.1% of thetotal myofibers. We did not observe any miniDys–GFP-positive fibers in DPMs from transplanted animalsanalyzed 2 or 4 months after transplantation. At 6 monthspost-transplantation, up to two miniDys–GFPþ myofiberswere detected in cryosections of the DPMs in three differentrecipients. This observation was consistent with the relativelylow levels of miniDys–GFP transcripts in the DPMs of themdx transplant recipients (Fig. 5C). No miniDys–GFPþ

myofibers were detected in the heart or liver of any recipients(data not shown).

Muscle damage can greatly facilitate incorporation ofBM cells into myofibers (31,48,50). The skeletal muscles ofmdx mice undergo continuous cycles of myofiber degenerationand regeneration (51,52). When mdx skeletal musclesare subjected to X-irradiation (as is done prior to BM

transplantation), progressive myofiber loss accompanied byincreasing fibrosis has been observed (53). At 6 months post-transplantation, all of our lethally irradiated mdx4Cv mouseBM transplant recipients were comparatively thin and weakand their harvested skeletal muscles appeared somewhatmore dystrophic compared with control mdx4Cv mice (datanot shown). We therefore asked whether more miniDys–GFP-expressing fibers were detected in mdx4Cv recipientssacrificed at increasingly longer time points after transplan-tation. Figure 5E shows a trend of increasing numbers ofminiDys–GFPþ myofibers in quadriceps of recipientanimals at greater intervals following BM transplantation,although this was not statistically significant. Unfortunately,we were not able to examine BM recipients at longer timepoints than 6 months due to a high rate of mouse mortalityin recipients. The highest average number of miniDys–GFPþ myofibers per 10 mm cryosection of quadricepsmuscles was only 10 at 6 months post-transplantation.Taken together, these data indicated that BM cells incorpor-ated into dystrophic mdx4Cv skeletal myofibers at a low fre-quency, conferring dystrophin expression, and the number ofdonor-derived dystrophin-positive fibers increased with time.This relatively low level of engraftment is similar to thatreported previously by others using WT BM donor cells(28,48,49).

DISCUSSION

Stem cell transplantation has emerged as a promisingapproach to developing a treatment for the muscular dystro-phies (54,55). The primary limitation for this approach is agreat need for improved efficiencies of cell transplantationinto dystrophic models. Consequently, research has focussedon identifying better cell sources for transplantation, under-standing the molecular mechanisms by which transplantedcells can home to and engraft in muscle and developingmethods for genetically correcting autologous stem cellsfor therapy (ex vivo gene therapy). We engineered aminiDys–GFP fusion gene to facilitate these studies byenabling easier tracing of donor cells that overexpress ahighly functional dystrophin protein. The miniDys–GFPcDNA is only 5.7 kb, such that it can be carried by lenti-viral vectors, which are a promising gene transfer vehiclefor genetic modification of mdx progenitor cells(12,13,56). Furthermore, we showed that at least threetypes of stem cells, myoblasts, MAPCs and BM cellscould be isolated from the transgenic mdx mice and giverise to miniDys-positive myofibers when transplanted intomdx4Cv mice.

The advantage of using donor cells that express a func-tional, traceable dystrophin is that engrafted myofibers willbe protected from necrosis and display a selective advantagerelative to dystrophin-negative myofibers that express a repor-ter gene such as lacZ. MiniDys-positive myofibers are easilydistinguished from revertant dystrophin-positive myofibersby the eGFP moiety. The success of this strategy requiresthat the miniDys–GFP fusion protein be highly functional,yet small enough to be carried by lentiviral vectors. The con-struct we used here had not been previously tested, so it was


critical to ensure that the miniDys fusion protein was capableof rescuing the mdx mouse muscle phenotype. Our resultsshow that this mini-dystrophin, which lacks two-thirds of thecentral rod domain as well as the CT domain, was almostfully functional. The only abnormality observed was a slightreduction in force generation in the transgenic mouse EDLmuscle. All other transgenic mouse muscles examineddisplayed normal force development, and the transgenicEDL displayed normal mass, histology and resistance tocontraction-induced injury. The reasons for this force dropare not clear, but do not appear to reflect functionaldeficiencies in the construct because the other muscles werenormal. Although dystrophins carrying either the rod or theCT domain truncations have been tested previously in trans-genic mice, the present study is the first to test a dystrophinlacking two separate domains in transgenic mdx mice(17,22). This study is also the first report of a detailed charac-terization of the physiological and mechanical properties ofthe 4Cv strain of mdx mice. The mdx4Cv model for DMDhas several potential advantages for studying stem cell andgene therapies. First, it displays fewer revertant fibers thando mdx mouse muscles (37). Secondly, the mdx4Cv mutationeliminates expression of several non-muscle isoforms of dys-trophin, potentially enabling a cleaner analysis of immuneresponses against exogenous dystrophin (57). The mechanicalproperties of muscles of mdx4Cv mice (Fig. 4) were quitesimilar to those observed previously for muscles of mdxmice (3,58).

A previous study also used eGFP as a marker to trace dys-trophin expression (59). This group fused the GFP-codingsequence with the N-terminus of dystrophin. Althoughin vivo DNA transfection showed that GFP–dystrophinfusion protein correctly localized on the sarcolemma of myo-fibers, its overall function in mdx mice was not evaluated. Theconstruct utilized a full-length dystrophin cDNA, precludingits use in lentiviral vectors. The ability to generate stablyexpressed and proper localizing GFP fusion constructs atboth the N-terminal and CT regions of dystrophin highlightsthe unusual ability of dystrophin to tolerate multiple structuralmodifications without adversely affecting its functionalcapacity. In particular, the ability to replace the CT domainof dystrophin with GFP suggests that a variety of other inser-tions in this region may also be compatible with nearly fulldystrophin function.

In the BM transplantation studies using the miniDys–GFP/mdx4Cv transgenic mice as donors, we observed that thenumber of donor-derived dystrophin-positive fibers in skeletalmuscles of mdx4Cv recipients increased with time (Fig. 5E).This increase likely reflects the continuous cycles of myofibernecrosis and regeneration that characterize mdx muscles,providing a niche for ongoing recruitment of BM cells intodystrophic myofibers. Several groups have reported thatmuscle injury greatly enhances attraction and engraftment ofhematopoietic stem cells in muscles of WT mice (31,48,50).The observations presented here are in agreement with thosefindings. Surprisingly, we also noticed that the DPM, whichis the most dystrophic striated muscle in mdx mice (60), dis-played a very low level of miniDys–GFP mRNA (Fig. 5C)and no detectable donor-derived myofibers even afterimmunostaining for GFP at 2 or 4 months post-transplantation.

At 6 months post-transplantation, the longest time point in thisstudy, recipient skeletal muscles were extremely dystrophicand the number of miniDys–GFPþ fibers remained low.Similar observations were reported when WT mice wereused as donors for whole BM transplantation to mdx4Cv mice(49). Thus, although a dystrophic microenvironment canenhance donor cell incorporation into muscle, it is notnearly sufficient to lead to a therapeutically significant levelof engraftment.

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

The mechanisms by which BM stem cells incorporate intomyofibers remain unclear. LaBarge and Blau (50) reportedthat donor hematopoietic cells can form myofibers via thecanonical myogenic pathway. Other groups have suggestedthat the low frequency of donor-derived myofibers occurredvia fusion of myeloid cells or BM stromal cells with myofibers(48,62). This latter study did not find any donor-derived myo-blast colonies when mononuclear muscle cells were culturedin vitro (48), nor we have been able to isolate such coloniesfrom our mdx4Cv BM transplant recipients (data not shown).However, high levels of infiltrating macrophages and neutro-phils are found in injured or dystrophic muscles (63,64) andmyeloid cells have been observed to fuse with myofibersin vivo (48). Interestingly, IL-4 has been shown to facilitatecell–cell fusion of myoblasts to myotubes as well as macro-phage fusion to form giant cells (65–67). In addition, thefusion of BM cells to hepatocytes, cardiomyocytes andPurkinje neurons after BM transplantation has been reportedby several groups (68–71). Further studies to understand theunderlying mechanisms of myofiber engraftment are clearlyneeded to improve the efficiency of stem cell strategies forDMD, and such studies may be aided by the reagentsdescribed in this paper.

In summary, the miniDys-GFP gene encodes a small andeasily traceable, but highly functional protein. Transgenic/mdx4Cv mice overexpress this fusion protein exclusively inskeletal muscles, providing a rich source of cells with whichto develop and optimize cell transplant therapies for DMD.Transplanted cells can be readily distinguished from revertantmdx myofibers, and the expressed dystrophin provides func-tional rescue and selection for engrafted myofibers. Thesame miniDys–GFP expression cassette is small enough tobe readily cloned into lentiviral vectors, making it a poten-tially useful construct with which to study approaches for exvivo gene therapy of DMD using autologous cell transplants.A considerable increase in our understanding of the bestcell source for transplants, optimal methods to isolate or pro-pagate those cells and the mechanisms underlying cell homingand engraftment in muscle are needed before cell therapiescan be useful for the muscular dystrophies. The miniDys–GFP gene and transgenic mice may help facilitate thesestudies.


MATERIALS AND METHODS

miniDys–GFP/mdx4Cv transgenic mice

A miniDys–GFP fusion gene was generated in several cloningsteps. We first replaced TAGGAA (underlined nucleotides arethe stop codon of dystrophin gene) with TCTAGA (XbaI site)in pSV40pAD71-78 (17) using the QuikChangew XL Site-Directed Mutagenesis Kit (Stratagene) (forward primer:50-GGAAACTGACACAATTCTAGAAGTCTTTTCCAC-30;reverse primer: 50-GTGGAAAAGACTTCTAGAATTGTGTCAGTTTCC-30). A PCR-amplified XbaI-flanked eGFP-coding sequence without its translational initiation region(forward primer: 50-GCTCTAGAGGTGAGCAAGGGCGAGGAG-30; reverse primer: 50-CCTCTAGAATTCCGGCCGCTTT AC-30) was then inserted in frame to generatepSV40pAD71-78-GFP. A 2222-bp HindIII fragment fromthis plasmid was used to replace the 1478-bp HindIIIfragment in the pCK6-DR4-R23D71-78 (17) to generatepCK6-DR4-R23D71-78-GFP. We then replaced the 2546-bpNsiI–BspEI fragment in the pCK6-DR4-R23D71-78-GFPwith a 3929 bp NsiI–BspEI fragment from pBSX-DH2-R19(17) to generate pCK6DH2-R19D71-78-eGFP (CK6-miniDys–eGFP). Finally, a 5128-bp EagI fragment containingDH2-R19D71-78-eGFP (miniDys–eGFP) in pCK6DH2-R19D71-78-GFP was inserted into pHSAvpSV40p expressionvector (22) at the NotI site to generate pHSA-DH2-R19D71-78-GFP-pA. The excised miniDys–GFP expressioncassette was injected into B6C3F1 hybrid fertilized eggs(Taconic), and F0 mice were screened for the transgene byPCR. Four positive F0 mice were backcrossed onto theB6Ros.Cg-Dmdmdx-4Cv (‘mdx4Cv’) background. Most studiesfocussed on the line 18916 that had the most uniform transgeneexpression levels in skeletal muscles.

Morphological assays

Quadriceps, TA, EDL and DPM muscles were removedfrom mice, embedded in O.C.T. medium (Tissue-Tek),frozen in liquid nitrogen and cut into 5 mm sections. After fix-ation in 3.7% formaldehyde, sections were stained inhematoxylin and eosin-phloxine. Stained sections wereimaged with a Nikon E1000 microscope connected to aSpot-2 CCD camera. To determine the percentage of myofi-bers containing central nuclei, the number of muscle fiberswith centrally located nuclei was divided by the totalnumber of muscle fibers. Evans blue assays were as describedpreviously (72).

Immunofluorescence analysis

As described earlier, 5 mm cryosections of various tissueswere prepared. Immunofluorescence detection was performedwith primary antibodies recognizing the followingproteins: the N-terminal domain of dystrophin (41), greenfluorescence protein (Molecular Probes), b-dystroglycan anda-, b-, g-sarcoglycan (Novacastra Laboratories, Ltd) anda1-syntrophin (73). After incubation with primary antibodies,the cryosections were incubated with a goat anti-rabbit or goatanti-mouse secondary antibody conjugated with Alexa 594(Molecular Probes). Images were collected on a Nikon

E1000 microscope under identical conditions using a Spot-2CCD camera.

Measurement of contractile properties

Contractile properties were measured in vitro on EDL andsoleus muscles and DPM strips obtained from7–10-month-old WT C57BL/6, mdx4Cv and miniDys–GFP/mdx4Cv transgenic male mice by methods described previously(74). An exception was the control data on DPM strips thatwere obtained from 17–18-month-old female WT mice.Control data on DPM strips of WT mice do not differamong mice from 6 to 24 months of age and between malesand females (75). EDL and soleus muscles were evaluatedfrom only one leg. In all, seven EDL, seven soleus and 16DPM muscles were examined from the mdx4Cv mice, sevenEDL, eight soleus and 16 DPM muscles from the transgenicmice and eight EDL, eight soleus and 15 DPM from WTmice. Mice were anesthetized with an intraperitoneal injectionof 1.3% avertin (0.015 ml/g body mass) with supplementalinjections as required to prevent response to tactile stimuli.EDL and soleus muscles were isolated and proximal anddistal tendons were tied firmly with silk (5.0) suture. EntireDPM muscles were excised. Muscles were removed fromthe mouse and submerged in a bath that contained bufferedmammalian Ringer’s solution (75). The intact fiber lengthsof DPM strips extended from the central tendon to the rib.Ties were placed firmly around both the central tendon andthe rib. For each of the three muscles, with the muscle in abath and platinum electrodes placed on either side, one endof the tendon was tied to a force transducer (model BG-50,Kulite Semiconductor Products) and the other tendon to thelever arm of a servomotor (model 305B Aurora Scientific,Richmond Hill, ON, Canada). Muscles were stimulateddirectly with a pulse duration of 2 ms. With the muscle atresting length, the voltage was increased to produce amaximum twitch force and muscle length was adjusted tooptimum length (Lo) for force development. With the musclelength set at Lo, the stimulation frequency was increaseduntil the development of force plateaued at the maximum iso-metric tetanic force (Po). Stimulation durations were of300 ms for EDL muscles and 900 ms for soleus and DPMmuscles. Po was usually occurred at a frequency of�180 Hz for EDL and �150 Hz for soleus and DPMmuscles. The susceptibility of muscles to contraction-inducedinjury was assessed by two lengthening contractions. Themuscles were set at Lo, activated maximally, and thenstretched through a strain of 30% at velocity of 1 Lf/s andthen returned at the same velocity to Lo, relaxed for a 10 srecovery period and then exposed to a second stretch of30%. The muscles were then allowed to recover for 1 minbefore the maximum force was measured. Muscles wereremoved from the bath, trimmed, blotted, weighed and sub-sequently frozen in isopentane cooled by dry ice. The totalfiber cross-sectional area (CSA, cm2) was calculated basedon the measurements of optimal muscle length (mm),muscle mass (mg), a muscle density of 1.06 g/cm2 and anLf/Lo ratio of 0.44 for the EDL, 0.71 for the soleus and 1 forthe DPM muscles (74). The specific Po (kN/m2) was deter-mined by dividing Po (kN) by CSA (m2). The force deficit


produced by the lengthening contraction protocol was assessedby expressing the Po (mN) measured after the two lengtheningcontraction protocol as a percentage of the Po (mN) beforeinjury.

Western analysis

Skeletal muscle microsomes from 4-month-old C57BL/6,mdx4Cv and miniDys–GFP/mdx4Cv transgenic mice (line18916) were prepared as described previously (76) and usedfor western analysis. The final microsome pellet was resus-pended in 0.3 M sucrose and 20 mM Tris-maleate (pH 7.0).Protein concentrations of each sample were measured by theBradford assay (77), and equal protein loading was verifiedby SDS–PAGE. KCl-washed microsomes were electrophor-etically separated on 4–12% gradient SDS–PAGE polyacryl-amide gels (BioRad) and analyzed by western blot using theprimary antibodies used for immunostaining, described earlier.

Detection of MHC loci

The MHC loci in C57BL/6 and C3 mouse strains can bedistinguished by PCR amplification of two markers(D17Mit10 and D17Mit51) around the MHC locus (http://jaxmice.jax.org). F2 mice from line 18916 were examinedby PCR and males carrying the two C57BL/6 markers werechosen for further backcrossing with femaleB6Ros.Cg-Dmdmdx-4Cv mice (‘mdx4Cv’) (Jackson Laboratory,Bar Harbor, ME, USA) for seven generations prior to usingthem for cell transplantation into mdx4Cv mouse recipients.

Preparation of primary myoblasts and intramusculartransplantation

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 1�phosphate-bufferedsaline (PBS) (pH 7.2) with 1 mM CaCl2 at 378C for 45 min.The digestion was stopped by adding F10 (GIBCO BRL)with 15% horse serum (Atlanta Biologicals). Digestedmuscle was filtered through 70 mm and then 40 mm nylonfilters (BD Falcon). Mononuclear cells were cultured ingelatin-coated plates (0.67%) with F10C (GIBCO BRL) sup-plemented with 15% horse serum (Atlanta Biologicals) and5 ng/ml basic FGF2 (R & D system) for 48 h to enrich formyoblasts. Cells were trypsinized, washed once and resus-pended in 1�PBS (pH 7.2). For transplantation, 2-month-oldmdx4Cv mice were anesthetized via sustained inhalation ofisoflurane in medical oxygen at a concentration sufficient toeliminate response to tactile stimuli. Their TA muscles weresurgically exposed. Forty microliters containing 5 � 105

cells suspension was loaded in 1 cc syringes and injected long-itudinally into each TA muscle. After injection, the skinincision was closed with cyanoacrylate adhesive (NexabandS/C, World Precision Instruments, Sarasota, FL, USA).

Isolation of muscle MAPCs

Limb muscles of miniDys–GFP/mdx4Cv transgenic mice wereexcised and mononuclear muscle cells were prepared as

described above. We immediately cultured these cells inmouse MAPC media as described to obtain muscle MAPCs(47). Before intramuscular injection, the multipotent differen-tiation capacity of isolated MAPCs was examined by in vitrodifferentiation assays. The MAPCs were trypsinized, washedand resuspended in 1�PBS (pH 7.2). As described above,7.5 � 104 MAPCs were intramuscularly injected into eachTA muscle of 2-month-old mdx4Cv mice.

Intravenous transplantation of whole BM cells

Whole BM cells were sterilely collected by a slightly modifiedmethod described by LaBarge and Blau (50). Briefly,6–8-week-old F5 miniDys–GFP/mdx4Cv transgenic micewere sacrificed by cervical dislocation, briefly immersed in75% ethanol and had their skin peeled back from a midline,circumferential incision. The femurs were dissociated andthe muscles around them removed, and the marrow cavitieswere flushed with a 26-gage needle containing ice-coldIscove’s DMD medium (GIBCO BRL) with 2% fetal bovineserum (FBS). Marrow cells were further dissociated by tritur-ating through the 26-gage needles. The resulting single-cellsuspension was spun at 300g for 10 min, and the pellet wasresuspended in ice-cold Iscove’s DMD with 2% FBS to 10million nucleated cells per ml. Two-month-old femalemdx4Cv mice were then lethally irradiated (1100 rad) andimmediately injected with 200 ml of 2 million nucleatedcells 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 isolatedwith an RNease Mini Kit (Qiagen), respectively, accordingto manufacturer’s instructions. Primer pairs (forward primer:50-AGCCGAGAGTAGCAGTTGTAGC-30; reverse primer:50-TTACATTTTTGACCTGCCAGTG-30) were designed(Fig. 1A) to specifically amplify a 470 bp fragment from theminiDys–GFP mRNA, and another primer pair (forwardprimer: 50-TGTGACGTTGACATCCGTAAAG-30; reverseprimer: 50-AAACGCAGCTCAGTAACAGTCC) was used toamplify a 300 bp fragment of b-actin mRNA. QIAGENOneStep RT–PCR kits were used to detect both miniDys–eGFP and b-actin mRNAs. For amplifying b-actin mRNAfragments (loading control), 25 ng total RNA from eachsample was used, whereas for miniDys–eGFP transcripts,250 ng total RNA was used.

ACKNOWLEDGEMENTS

We thank X. Ye and G. Stamatoyannopoulos for the gener-ation of MiniDys–GFP transgenic mice and J. Yan,M. Weinreich, J.C. Angello, C.A. Blau and S.D. Hauschkafor advice and technical assistance. We also thank allmembers in the Chamberlain lab for helpful discussions andadvice. These studies were supported by grants from theNational Institutes of Health (P01 AG015434, P01 NS46788and 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 DystrophyAssociation (USA).

Conflict of Interest statement. None declared.

REFERENCES

1. Emery, A.E.H. (ed.), (2003) The Muscular Dystrophies, 2nd edn. OxfordUniversity Press Inc., New York, Oxford.

2. Abmayr, S. and Chamberlain, J.S. (2004) The structure and function ofdystrophin. In Winder, S. (ed.), The Molecular Mechanisms in MuscularDystrophy. Landes Biosciences, Georgetown.

3. Lynch, G.S., Hinkle, R.T., Chamberlain, J.S., Brooks, S.V. andFaulkner, J.A. (2001) Force and power output of fast and slowskeletal muscles from mdx mice 6–28 months old. J. Physiol., 535,591–600.

4. Lynch, G.S., Rafael, J.A., Chamberlain, J.S. and Faulkner, J.A. (2000)Contraction-induced injury to single permeabilized muscle fibers frommdx, transgenic mdx, and control mice. Am. J. Physiol. Cell Physiol., 279,C1290–C1294.

5. Petrof, B.J., Shrager, J.B., Stedman, H.H., Kelly, A.M. and Sweeney, H.L.(1993) Dystrophin protects the sarcolemma from stresses developedduring muscle contraction. Proc. Natl Acad. Sci. USA, 90, 3710–3714.

6. Rando, T.A. (2001) The dystrophin–glycoprotein complex, cellularsignaling, and the regulation of cell survival in the muscular dystrophies.Muscle Nerve, 24, 1575–1594.

7. Ohlendieck, K. and Campbell, K.P. (1991) Dystrophin-associated proteinsare greatly reduced in skeletal muscle from mdx mice. J. Cell Biol., 115,1685–1694.

8. Chamberlain, J.S. (2002) Gene therapy of muscular dystrophy. Hum. Mol.Genet., 11, 2355–2362.

9. Gregorevic, P. and Chamberlain, J.S. (2003) Gene therapy for musculardystrophy—a review ofpromising progress. Expert Opin. Biol. Ther., 3,803–814.

10. Voisin, V. and de la Porte, S. (2004) Therapeutic strategies for Duchenneand Becker dystrophies. Int. Rev. Cytol., 240, 1–30.

11. Bogdanovich, S., Perkins, K.J., Krag, T.O. and Khurana, T.S. (2004)Therapeutics for Duchenne muscular dystrophy: current approaches andfuture directions. J. Mol. Med., 82, 102–115.

12. Bachrach, E., Li, S., Perez, A.L., Schienda, J., Liadaki, K., Volinski, J.,Flint, A., Chamberlain, J. and Kunkel, L.M. (2004) Systemic delivery ofhuman microdystrophin to regenerating mouse dystrophic muscle bymuscle progenitor cells. Proc. Natl Acad. Sci. USA, 101, 3581–3586.

13. Li, S., Kimura, E., Fall, B.M., Reyes, M., Angello, J.C., Welikson, R.,Hauschka, S.D. and Chamberlain, J.S. (2005) Stable transduction ofmyogenic cells with lentiviral vectors expressing a minidystrophin. GeneTher., 12, 1099–1108.

14. Ailles, L., Schmidt, M., Santoni de Sio, F.R., Glimm, H., Cavalieri, S.,Bruno, S., Piacibello, W., Von Kalle, C. and Naldini, L. (2002) Molecularevidence of lentiviral vector-mediated gene transfer into humanself-renewing, multi-potent, long-term NOD/SCID repopulatinghematopoietic cells. Mol. Ther., 6, 615–626.

15. Sampaolesi, M., Torrente, Y., Innocenzi, A., Tonlorenzi, R.,D’Antona, G., Pellegrino, M.A., Barresi, R., Bresolin, N., DeAngelis, M.G., Campbell, K.P. et al. (2003) Cell therapy ofalpha-sarcoglycan null dystrophic mice through intra-arterial delivery ofmesoangioblasts. Science, 301, 487–492.

16. Wang, B., Li, J. and Xiao, X. (2000) Adeno-associated virus vectorcarrying human minidystrophin genes effectively ameliorates musculardystrophy in mdx mouse model. Proc. Natl Acad. Sci. USA, 97,13714–13719.

17. Harper, S.Q., Hauser, M.A., DelloRusso, C., Duan, D., Crawford, R.W.,Phelps, S.F., Harper, H.A., Robinson, A.S., Engelhardt, J.F., Brooks, S.V.et al. (2002) Modular flexibility of dystrophin: implications for genetherapy of Duchenne muscular dystrophy. Nat. Med., 8, 253–261.

18. Sakamoto, M., Yuasa, K., Yoshimura, M., Yokota, T., Ikemoto, T.,Suzuki, M., Dickson, G., Miyagoe-Suzuki, Y. and Takeda, S. (2002)Micro-dystrophin cDNA ameliorates dystrophic phenotypes whenintroduced into mdx mice as a transgene. Biochem. Biophys. Res.Commun., 293, 1265–1272.

19. Watchko, J., O’Day, T., Wang, B., Zhou, L., Tang, Y., Li, J. and Xiao, X.(2002) Adeno-associated virus vector-mediated minidystrophin gene

therapy improves dystrophic muscle contractile function in mdx mice.Hum. Gene Ther., 13, 1451–1460.

20. Gregorevic, P., Blankinship, M.J., Allen, J.M., Crawford, R.W.,Meuse, L., Miller, D.G., Russell, D.W. and Chamberlain, J.S. (2004)Systemic delivery of genes to striated muscles using adeno-associatedviral vectors. Nat. Med., 10, 828–834.

21. Abmayr, S., Gregorevic, P., Allen, J.M. and Chamberlain, J.S.(2005) Phenotypic improvement of dystrophic muscles by rAAV/microdystrophin vectors is augmented by Igf1 codelivery. Mol. Ther., 12,441–450.

22. Crawford, G.E., Faulkner, J.A., Crosbie, R.H., Campbell, K.P.,Froehner, S.C. and Chamberlain, J.S. (2000) Assembly of thedystrophin-associated protein complex does not require the dystrophinCOOH-terminal domain. J. Cell Biol., 150, 1399–1410.

23. Yoshida, M., Hama, H., Ishikawa-Sakurai, M., Imamura, M., Mizuno, Y.,Araishi, K., Wakabayashi-Takai, E., Noguchi, S., Sasaoka, T. andOzawa, E. (2000) Biochemical evidence for association of dystrobrevinwith the sarcoglycan–sarcospan complex as a basis for understandingsarcoglycanopathy. Hum. Mol. Genet., 9, 1033–1040.

24. Balasubramanian, S., Fung, E.T. and Huganir, R.L. (1998)Characterization of the tyrosine phosphorylation and distribution ofdystrobrevin isoforms. FEBS Lett., 432, 133–140.

25. Sadoulet-Puccio, H.M., Rajala, M. and Kunkel, L.M. (1997) Dystrobrevinand dystrophin: an interaction through coiled-coil motifs. Proc. Natl Acad.Sci. USA, 94, 12413–12418.

26. Peters, M.F., Sadoulet-Puccio, H.M., Grady, M.R., Kramarcy, N.R.,Kunkel, L.M., Sanes, J.R., Sealock, R. and Froehner, S.C. (1998)Differential membrane localization and intermolecular associations ofalpha-dystrobrevin isoforms in skeletal muscle. J. Cell. Biol., 142,1269–1278.

27. Ishikawa-Sakurai, M., Yoshida, M., Imamura, M., Davies, K.E. andOzawa, E. (2004) ZZ domain is essentially required for the physiologicalbinding of dystrophin and utrophin to beta-dystroglycan. Hum. Mol.Genet., 13, 693–702.

28. Gussoni, E., Soneoka, Y., Strickland, C.D., Buzney, E.A., Khan, M.K.,Flint, A.F., Kunkel, L.M. and Mulligan, R.C. (1999) Dystrophinexpression in the mdx mouse restored by stem cell transplantation.Nature, 401, 390–394.

29. Montanaro, F., Liadaki, K., Volinski, J., Flint, A. and Kunkel, L.M. (2003)Skeletal muscle engraftment potential of adult mouse skin side populationcells. Proc. Natl Acad. Sci. USA, 100, 9336–9341.

30. De Bari, C., Dell’Accio, F., Vandenabeele, F., Vermeesch, J.R.,Raymackers, J.M. and Luyten, F.P. (2003) Skeletal muscle repair by adulthuman mesenchymal stem cells from synovial membrane. J. Cell Biol.,160, 909–918.

31. Ferrari, G., Cusella-De Angelis, G., Coletta, M., Paolucci, E.,Stornaiuolo, A., Cossu, G. and Mavilio, F. (1998) Muscle regeneration bybone marrow-derived myogenic progenitors. Science, 279, 1528–1530.

32. Bittner, R.E., Schofer, C., Weipoltshammer, K., Ivanova, S., Streubel, B.,Hauser, E., Freilinger, M., Hoger, H., Elbe-Burger, A. and Wachtler, F.(1999) Recruitment of bone-marrow-derived cells by skeletal and cardiacmuscle in adult dystrophic mdx mice. Anat. Embryol. (Berl.), 199,391–396.

33. Valentine, B.A., Winand, N.J., Pradhan, D., Moise, N.S., de Lahunta, A.,Kornegay, J.N. and Cooper, B.J. (1992) Canine X-linked musculardystrophy as an animal model of Duchenne muscular dystrophy: a review.Am. J. Med. Genet., 42, 352–356.

34. Wilton, S.D., Dye, D.E., Blechynden, L.M. and Laing, N.G. (1997)Revertant fibres: a possible genetic therapy for Duchenne musculardystrophy? Neuromuscul. Disord., 7, 329–335.

35. Sicinski, P., Geng, Y., Ryder-Cook, A.S., Barnard, E.A., Darlison, M.G.and Barnard, P.J. (1989) The molecular basis of muscular dystrophy in themdx mouse: a point mutation. Science, 244, 1578–1580.

36. Dell’Agnola, C., Wang, Z., Storb, R., Tapscott, S.J., Kuhr, C.S.,Hauschka, S.D., Lee, R.S., Sale, G.E., Zellmer, E., Gisburne, S. et al.(2004) Hematopoietic stem cell transplantation does not restoredystrophin expression in Duchenne muscular dystrophy dogs.Blood, 104, 4311–4318.

37. Danko, I., Chapman, V. and Wolff, J.A. (1992) The frequency ofrevertants in mdx mouse genetic models for Duchenne musculardystrophy. Pediatr. Res., 32, 128–131.

38. Lu, Q.L., Morris, G.E., Wilton, S.D., Ly, T., Artem’yeva, O.V., Strong, P.and Partridge, T.A. (2000) Massive idiosyncratic exon skipping corrects


the nonsense mutation in dystrophic mouse muscle and producesfunctional revertant fibers by clonal expansion. J. Cell Biol., 148,985–996.

39. Brennan, K.J. and Hardeman, E.C. (1993) Quantitative analysis of thehuman alpha-skeletal actin gene in transgenic mice. J. Biol. Chem., 268,719–725.

40. Kozak, M. (1986) Point mutations define a sequence flanking the AUGinitiator codon that modulates translation by eukaryotic ribosomes. Cell,44, 283–292.

41. Rafael, J.A., Cox, G.A., Corrado, K., Jung, D., Campbell, K.P. andChamberlain, J.S. (1996) Forced expression of dystrophin deletionconstructs reveals structure-function correlations. J. Cell Biol., 134,93–102.

42. Torres, L.F. and Duchen, L.W. (1987) The mutant mdx: inheritedmyopathy in the mouse. Morphological studies of nerves, muscles andend-plates. Brain, 110 (Pt 2), 269–299.

43. Bockhold, K.J., Rosenblatt, J.D. and Partridge, T.A. (1998) Aging normaland dystrophic mouse muscle: analysis of myogenicity in cultures ofliving single fibers. Muscle Nerve, 21, 173–183.

44. Matsuda, R., Nishikawa, A. and Tanaka, H. (1995) Visualization ofdystrophic muscle fibers in mdx mouse by vital staining with Evans blue:evidence of apoptosis in dystrophin-deficient muscle. J. Biochem. (Tokyo),118, 959–964.

45. Judge, L., Haraguchi, M. and Chamberlain, J.S. (2006) Dissecting thesignaling and mechanical functions of the dystrophin–glycoproteincomplex. J. Cell Sci., in press.

46. Lai, Y., Yue, Y., Liu, M., Ghosh, A., Engelhardt, J.F., Chamberlain, J.S.and Duan, D. (2005) Efficient in vivo gene expression by trans-splicingadeno-associated viral vectors. Nat. Biotechnol., 23, 1435–1439.

47. Jiang, Y., Vaessen, B., Lenvik, T., Blackstad, M., Reyes, M. andVerfaillie, C.M. (2002) Multipotent progenitor cells can be isolated frompostnatal murine bone marrow, muscle, and brain. Exp. Hematol., 30,896–904.

48. Camargo, F.D., Green, R., Capetenaki, Y., Jackson, K.A. andGoodell, M.A. (2003) Single hematopoietic stem cells generate skeletalmuscle through myeloid intermediates. Nat. Med., 9, 1520–1527.

49. Ferrari, G., Stornaiuolo, A. and Mavilio, F. (2001) Failure to correctmurine muscular dystrophy. Nature, 411, 1014–1015.

50. LaBarge, M.A. and Blau, H.M. (2002) Biological progression from adultbone marrow to mononucleate muscle stem cell to multinucleate musclefiber in response to injury. Cell, 111, 589–601.

51. Coulton, G.R., Morgan, J.E., Partridge, T.A. and Sloper, J.C. (1988) Themdx mouse skeletal muscle myopathy: I. A histological, morphometricand biochemical investigation. Neuropathol. Appl. Neurobiol., 14, 53–70.

52. McGeachie, J.K., Grounds, M.D., Partridge, T.A. and Morgan, J.E. (1993)Age-related changes in replication of myogenic cells in mdxmice: quantitative autoradiographic studies. J. Neurol. Sci., 119,169–179.

53. Wakeford, S., Watt, D.J. and Partridge, T.A. (1991) X-irradiationimproves mdx mouse muscle as a model of myofiber loss in DMD. Muscle

Nerve, 14, 42–50.54. Chakkalakal, J.V., Thompson, J., Parks, R.J. and Jasmin, B.J. (2005)

Molecular, cellular, and pharmacological therapies for Duchenne/Beckermuscular dystrophies. FASEB J., 19, 880–891.

55. Sohn, R.L. and Gussoni, E. (2004) Stem cell therapy for musculardystrophy. Expert Opin. Biol. Ther., 4, 1–9.

56. Kobinger, G.P., Louboutin, J.P., Barton, E.R., Sweeney, H.L. andWilson, J.M. (2003) Correction of the dystrophic phenotype byin vivo targeting of muscle progenitor cells. Hum. Gene Ther., 14,1441–1449.

57. Im, W.B., Phelps, S.F., Copen, E.H., Adams, E.G., Slightom, J.L. andChamberlain, J.S. (1996) Differential expression of dystrophin isoforms instrains of mdx mice with different mutations. Hum. Mol. Genet., 5,1149–1153.

58. Dellorusso, C., Crawford, R.W., Chamberlain, J.S. and Brooks, S.V.(2001) Tibialis anterior muscles in mdx mice are highly susceptible tocontraction-induced injury. J. Muscle Res. Cell Motil., 22, 467–475.

59. Chapdelaine, P., Moisset, P.A., Campeau, P., Asselin, I., Vilquin, J.T. andTremblay, J.P. (2000) Functional EGFP–dystrophin fusion proteins forgene therapy vector development. Protein Eng., 13, 611–615.

60. Stedman, H.H., Sweeney, H.L., Shrager, J.B., Maguire, H.C.,Panettieri, R.A., Petrof, B., Narusawa, M., Leferovich, J.M., Sladky, J.T.and Kelly, A.M. (1991) The mdx mouse diaphragm reproduces thedegenerative changes of Duchenne muscular dystrophy. Nature, 352,536–539.

61. Brazelton, T.R., Nystrom, M. and Blau, H.M. (2003) Significantdifferences among skeletal muscles in the incorporation of bonemarrow-derived cells. Dev. Biol., 262, 64–74.

62. Shi, D., Reinecke, H., Murry, C.E. and Torok-Storb, B. (2004) Myogenicfusion of human bone marrow stromal cells, but not hematopoietic cells.Blood, 104, 290–294.

63. Orimo, S., Hiyamuta, E., Arahata, K. and Sugita, H. (1991) Analysis ofinflammatory cells and complement C3 in bupivacaine-inducedmyonecrosis. Muscle Nerve, 14, 515–520.

64. Parrish, E.P., Cifuentes-Diaz, C., Li, Z.L., Vicart, P., Paulin, D.,Dreyfus, P.A., Peschanski, M., Harris, A.J. and Garcia, L. (1996)Targeting widespread sites of damage in dystrophic muscle: engraftedmacrophages as potential shuttles. Gene Ther., 3, 13–20.

65. McNally, A.K. and Anderson, J.M. (2002) Beta1 and beta2 integrinsmediate adhesion during macrophage fusion and multinucleated foreignbody giant cell formation. Am. J. Pathol., 160, 621–630.

66. McNally, A.K. and Anderson, J.M. (2003) Foreign body-typemultinucleated giant cell formation is potently induced byalpha-tocopherol and prevented by the diacylglycerol kinase inhibitorR59022. Am. J. Pathol., 163, 1147–1156.

67. Horsley, V., Jansen, K.M., Mills, S.T. and Pavlath, G.K. (2003) IL-4acts as a myoblast recruitment factor during mammalian muscle growth.Cell, 113, 483–494.

68. Alvarez-Dolado, M., Pardal, R., Garcia-Verdugo, J.M., Fike, J.R.,Lee, H.O., Pfeffer, K., Lois, C., Morrison, S.J. and Alvarez-Buylla, A.(2003) Fusion of bone-marrow-derived cells with Purkinje neurons,cardiomyocytes and hepatocytes. Nature, 425, 968–973.

69. Vassilopoulos, G., Wang, P.R. and Russell, D.W. (2003) Transplantedbone marrow regenerates liver by cell fusion. Nature, 422, 901–904.

70. Wang, X., Willenbring, H., Akkari, Y., Torimaru, Y., Foster, M.,Al-Dhalimy, M., Lagasse, E., Finegold, M., Olson, S. and Grompe, M.(2003) Cell fusion is the principal source of bone-marrow-derivedhepatocytes. Nature, 422, 897–901.

71. Weimann, J.M., Johansson, C.B., Trejo, A. and Blau, H.M. (2003) Stablereprogrammed heterokaryons form spontaneously in Purkinje neuronsafter bone marrow transplant. Nat. Cell Biol., 5, 959–966.

72. Straub, V., Rafael, J.A., Chamberlain, J.S. and Campbell, K.P. (1997)Animal models for muscular dystrophy show different patterns ofsarcolemmal disruption. J. Cell Biol., 139, 375–385.

73. Peters, M.F., Adams, M.E. and Froehner, S.C. (1997) Differentialassociation of syntrophin pairs with the dystrophin complex. J. Cell Biol.,138, 81–93.

74. Brooks, S.V. and Faulkner, J.A. (1988) Contractile properties of skeletalmuscles from young, adult and aged mice. J. Physiol., 404, 71–82.

75. Lynch, G.S., Rafael, J.A., Hinkle, R.T., Cole, N.M., Chamberlain, J.S. andFaulkner, J.A. (1997) Contractile properties of diaphragm musclesegments from old mdx and old transgenic mdx mice. Am. J. Physiol.,272, C2063–C2068.

76. Ohlendieck, K., Ervasti, J.M., Snook, J.B. and Campbell, K.P. (1991)Dystrophin–glycoprotein complex is highly enriched in isolated skeletalmuscle sarcolemma. J. Cell Biol., 112, 135–148.

77. Bradford, M.M. (1976) A rapid and sensitive method for the quantitationof microgram quantities of protein utilizing the principle of protein-dyebinding. Anal. Biochem., 72, 248–254.


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

Documents

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