Dystrophins carrying spectrin-like repeats 16 and 17 anchor nNOS to the sarcolemma and enhance exercise performance in a mouse model of muscular dystrophy

Sarcolemma-associated neuronal NOS (nNOS) plays a critical role in normal muscle physiology. In Duchenne muscular dystrophy (DMD), the loss of sarcolemmal nNOS leads to functional ischemia and muscle damage; however, the mechanism of nNOS subcellular localization remains incompletely understood. According to the prevailing model, nNOS is recruited to the sarcolemma by syntrophin, and in DMD this localization is altered. Intriguingly, the presence of syntrophin on the membrane does not always restore sarcolemmal nNOS. Thus, we wished to determine whether dystrophin functions in subcellular localization of nNOS and which regions may be necessary. Using in vivo transfection of dystrophin deletion constructs, we show that sarcolemmal targeting of nNOS was dependent on the spectrin-like repeats 16 and 17 (R16/17) within the rod domain. Treatment of mdx mice (a DMD model) with R16/17-containing synthetic dystrophin genes effectively ameliorated histological muscle pathology and improved muscle strength as well as exercise performance. Furthermore, sarcolemma-targeted nNOS attenuated α-adrenergic vasoconstriction in contracting muscle and improved muscle perfusion during exercise as measured by Doppler and microsphere circulation. In summary, we have identified the dystrophin spectrin-like repeats 16 and 17 as a novel scaffold for nNOS sarcolemmal targeting. These data suggest that muscular dystrophy gene therapies based on R16/17-containing dystrophins may yield better clinical outcomes than the current therapies.

Neuronal NOS (nNOS) plays critical roles in a variety of muscle activities, including contraction, regeneration, atrophy, glucose uptake, and blood perfusion (1). In normal skeletal muscle, nNOS is localized to the cytosolic surface of the sarcolemma. This position is critical for nNOS function. During muscle contraction, activated nNOS produces NO. The membrane location of nNOS allows ready diffusion of NO to the muscle vasculature. Consequently, vessels are dilated to meet the increased metabolic needs of contracting muscle (2, 3).

Duchenne muscular dystrophy (DMD) is caused by dystrophin deficiency (4). Despite the fact that dystrophin is missing in all myofibers, muscle disease does not occur simultaneously throughout the entire muscle. The characteristic early histopathology in DMD muscle closely resembles focal necrosis seen after functional ischemia (5, 6). The identification of nNOS as a component of the dystrophin-associated glycoprotein complex (DGC) provides a molecular explanation for the unique muscle lesions in DMD (7, 8). In the absence of dystrophin, the entire DGC (including nNOS) fails to assemble at the sarcolemma. Delocalization of nNOS impairs compensatory vasodilation during muscle contraction and subsequently leads to focal ischemia and necrosis (2, 3). For similar reasons, Duchenne patients are more easily fatigued than normal people (9, 10).

The mechanism(s) by which nNOS is recruited to the sarcolemma are not completely understood. According to the prevailing model, nNOS is brought to the sarcolemma by the scaffolding protein syntrophin. Several lines of evidence support this model. First, syntrophin itself is recruited to the sarcolemma by a syntrophin-binding motif in the dystrophin C-terminal domain (11–13). In the absence of the dystrophin C-terminal domain, syntrophin can still be recruited to the sarcolemma by dystrobrevin, possibly through the sarcoglycan complex (14–16). Second, syntrophin forms a complex with nNOS through a noncanonical interaction between PDZ (postsynaptic density 95, discs large, and zonula occludens–1) and PDZ domain (7, 8, 17, 18). Finally, genetic ablation of the syntrophin gene or the PDZ domain of the syntrophin gene eliminates sarcolemmal nNOS (19, 20).

Despite the compelling evidence, this model cannot explain all the existing data. Most intriguingly, the presence of syntrophin on the membrane does not always lead to sarcolemmal nNOS expression. This was first demonstrated in Becker muscular dystrophy (BMD), a milder allelic form of DMD (21). Synthetic mini-/microdystrophin genes that have been developed for DMD gene therapy also fail to anchor nNOS to the sarcolemma, although they are able to restore syntrophin (16, 22, 23). In addition, the dissociation between syntrophin and nNOS expression was observed in α-dystrobrevin–knockout mice (24). In these mice, the absence of α-dystrobrevin barely affects the syntrophin level, but nNOS is substantially reduced (24). Collectively, these results suggest a ternary interaction model for nNOS localization. Here, we hypothesized that in addition to a direct interaction between syntrophin and nNOS, one and/or multiple regions of dystrophin might have also contributed to nNOS localization by either directly interacting or indirectly stabilizing the syntrophin–nNOS complex.

To test this hypothesis, we generated a series of synthetic dystrophin genes and evaluated nNOS restoration by in vivo plasmid transfection, AAV-mediated gene transfer, and generation of a transgenic mouse model. After confirming dystrophin expression with a panel of epitope-specific antibodies, we examined nNOS expression by single and double immunostaining, in situ nNOS activity assay, and Western blot analysis. More than half of the dystrophin gene (from exon 3 to exon 52) has been implicated in nNOS localization (21). Surprisingly, we found that only a fairly small region in the middle of the dystrophin rod domain (R16 and R17, abbreviated as R16/17 herein) was required. Yeast two-hybrid assay results suggested that nNOS localization might be mediated by a coordinated effort of both R16/17 and the syntrophin PDZ domain, although the latter displayed a higher affinity for the nNOS PDZ domain. Based on these results, we developed several R16/17-containing mini- and microdystrophin genes as potential candidate genes for DMD/BMD gene therapy. In vivo studies in mouse models of DMD suggest that these newly developed candidate genes were as effective as the previously published mini- and microgenes in mitigating dystrophic histopathology and improving muscle force. In contrast to the previous candidate genes, the newly developed genes also restored sarcolemmal nNOS. In hemodynamic, tissue blood flow, and treadmill assays, R16/17-containing dystrophin significantly improved perfusion in contracting muscle and enhanced exercise performance.

Dystrophin recruits nNOS to the sarcolemma through the rod, not the C-terminal, domain. The full-length dystrophin gene can recruit nNOS to the sarcolemma, but the DH2–R19 minigene cannot (Figure 1 and Table 1) (23, 25). The DH2–R19 minigene has the same structure as the full-length gene, except for a large deletion in the rod domain. The deletion represents approximately 48% of the dystrophin coding sequence and includes 1 hinge (H2) and 16 spectrin-like repeats (from R4 to R19). To localize the nNOS recruiting domain, we gradually added back the deleted region to the human DH2–R19 minigene and evaluated the nNOS recruiting capacity (Figure 1, Table 1, and Supplemental Figures 1–3; supplemental material available online with this article; doi:10.1172/JCI36612DS1).

Figure 1

Sarcolemmal nNOS localization depends on the rod, but not the C-terminal, domain of dystrophin. (A) Schematic outline of the full-length and synthetic dystrophin constructs. “Yes” indicates that nNOS is recruited to the sarcolemma by the construct; “No” indicates that the construct cannot restore nNOS to the sarcolemma. Dotted boxes denote deleted regions. (B) The DH2–R15 minidystrophin anchors nNOS to the sarcolemma. Representative photomicrographs of dystrophin immunostaining, nNOS activity staining, and nNOS immunostaining in the DH2–R15 minigene plasmid–transfected mdx muscle. Dystrophin was revealed by epitope-specific antibodies (H1, R11, R16, R17, R18, and H3). The H1 antibody only recognizes human dystrophin (Hum Dys). Other dystrophin antibodies recognize both human and mouse dystrophin. Asterisks indicate a minigene-transfected myofiber; arrows, revertant myofibers. Scale bar: 20 μm. (C) Sarcolemmal nNOS localization does not require the C-terminal domain, nor is it dependent on the repeats adjacent to R16/17. Representative photomicrographs of human dystrophin/nNOS double immunostaining and nNOS activity staining on serial sections from the DH2–R15/DR18–19, DH2–R15/DC, and DR3–15/DR18–23/DC plasmid–transfected mdx muscles. Asterisks indicate dystrophin plasmid–transfected fibers; crosses, revertant myofibers. Scale bar: 50 μm.

Table 1

Sarcolemmal nNOS expression in dystrophin plasmid–transfected mdx muscle

Transfected myofibers were identified with a human dystrophin H1–specific antibody. nNOS expression was confirmed on serial sections by immunostaining and in situ nNOS activity assay. Sarcolemmal nNOS expression was not restored when R18/19 or R17/18/19 were inserted back into the DH2–R19 minigene (Table 1 and Supplemental Figures 2 and 3). However, restoring 4 repeats (R16/17/18/19) recovered sarcolemmal nNOS expression (Figure 1B, Table 1, and Supplemental Figure 2).

To determine whether all 4 added repeats contributed to nNOS restoration, we deleted R18/19 or R17/18/19 from the DH2–R15 minigene (Figure 1A). Removing R18/19 did not affect sarcolemmal nNOS expression (Figure 1C and Table 1). For unknown reasons, we were unable to detect DH2–R15/DR17–19 minigene expression in transfected mdx muscle (Figure 1A and Table 1).

Next, we evaluated the contribution of the dystrophin C-terminal domain. This domain has been suggested to indirectly facilitate nNOS recruitment by bringing syntrophin to the sarcolemma (11–13). However, when the C-terminal domain was deleted from the full-length gene, there was minimal impact on sarcolemmal nNOS expression (15). Consistent with this previous report, when the C-terminal domain was deleted from the DH2–R15 minigene, we did not detect any change in nNOS localization either (Figure 1C and Table 1). Together, these results suggest that the C-terminal domain is not required for nNOS recruitment in either the full-length gene or the abbreviated minigenes.

All 3 nNOS-recruiting minigenes (DH2–R15, DH2–R15/DR18–19, and DH2–R15/DC) shared 2 common features. They all contained a rod domain longer than that of the DH2–R19 minigene, and they all carried R16/17 with R16 flanked by R3. To determine whether the length of the rod domain and the configuration of the flanking repeats are important for nNOS recruitment, we generated the DR3–15/DR18–23/DC microgene. This microgene contains only 5 repeats. Furthermore, R16 is flanked by R2. Despite a much smaller rod domain and different flanking repeats, sarcolemmal nNOS expression was restored by the DR3–15/DR18–23/DC microgene (Figure 1C and Table 1).

In summary, our results suggest that R16/17 is crucial for dystrophin-mediated nNOS recruitment. Furthermore, the nNOS recruiting activity is independent of the C-terminal domain and independent of the neighboring repeats adjacent to R16/17.

nNOS localization requires both R16 and R17. The findings from the DH2–R16 construct suggest that the presence of R17 alone is not sufficient to recruit nNOS (Figure 1A, Table 1, Supplemental Figure 2, and Supplemental Figure 3D). Since we did not observe expression from the DH2–R15/DR17–19 minigene, it was not clear whether R16 by itself was sufficient for nNOS recruitment.

To address this issue, we generated 2 microdystrophin constructs. Both have 4 repeats in the rod domain. However, one contained only R16 (DR3–15/DR17–23/DC) and the other contained both R16 and R17 (DR2–15/DR18–23/DC) (Figure 2A). After packaging these constructs into adeno-associated virus 6 (AAV-6) vectors, we delivered them to the tibialis anterior (TA) muscles of 50-day-old mdx mice. At 30 days after AAV infection, we examined microgene and nNOS expression (Figure 2, A–C, and Supplemental Figure 4). We observed efficient transduction from both vectors (≥90%) (Figure 2, B and C). However, only the construct containing R16/17 (DR2–15/DR18–23/DC) restored sarcolemmal nNOS (Figure 2C and Supplemental Figure 4). These data strongly suggest that nNOS recruitment requires both R16 and R17 and that either one alone is not sufficient.

Figure 2

Both R16 and R17 are indispensable for sarcolemmal nNOS localization. (A) Schematic outline of the microdystrophin constructs. “Yes” indicates that nNOS is recruited to the sarcolemma by the construct; “No” indicates that nNOS is not recruited. Dotted boxes denote deleted regions. (B and C) Both R16 and R17 are required for sarcolemmal nNOS localization. Representative serial immunostaining and nNOS activity staining photomicrographs of mdx muscles infected with AV.CMV.DR3–15/DR17–23/DC (B) or AV.CMV.DR2–15/DR18–23/DC (C). Sarcolemmal nNOS localization was observed only in microdystrophin carrying both R16 and R17 (C) but not in microdystrophin carrying only R16 (B). Scale bars: 500 μm. See Supplemental Figure 4 for high-magnification images. (D) Representative yeast two-hybrid assay results (from 5 independent experiments) reveal interaction between R16/17 and the nNOS PDZ domain. Individual repeat R16 alone or R17 alone does not bind the nNOS PDZ domain. (E) Relative β-galactosidase activity from the quantitative yeast two-hybrid assay. n = 6 for each group.

R16/17 interacts with the nNOS PDZ domain. To further understand the contribution of R16/17 to nNOS recruiting, we performed a yeast two-hybrid assay (Figure 2, D and E). We cloned the nNOS PDZ domain into a binding construct. We also generated a series of the activation constructs carrying R16, R17, or R16/17. As a positive control, we also made an activation construct containing the syntrophin PDZ domain. Interaction with the nNOS PDZ domain was observed with R16/17 and the syntrophin PDZ domain, but not with R16 or R17 alone (Figure 2D). Similar results were obtained when the nNOS PDZ domain was cloned into the activation constructs, and other components (R16, R17, R16/17, or the syntrophin PDZ domain) were cloned into the binding constructs (data not shown). To compare the relative binding affinity for the nNOS PDZ domain, we performed a quantitative yeast two-hybrid assay (Figure 2E). Despite an absolute requirement of R16/17 for nNOS localization (Figures 1 and 2), the binding affinity between R16/17 and the nNOS PDZ domain was 5-fold lower than that between the syntrophin PDZ domain and the nNOS PDZ domain. Taken together, these results suggest that dystrophin R16/17 may assist sarcolemmal nNOS localization by interacting with the nNOS PDZ domain.

The R16/17-containing microgene ameliorates dystrophic pathology and improves muscle strength in mouse models for DMD. An important goal of this study is to improve current AAV-mediated DMD gene therapy. We have already demonstrated that an R16/17-containing microgene, DR2–15/DR18–23/DC, can restore sarcolemma nNOS (Figure 2 and Supplemental Figure 4). However, it is not clear whether this microgene can ameliorate dystrophic pathology. It is known that some microgenes cannot mitigate muscle disease (26).

To address this issue, we delivered AAV-6 AV.CMV.DR2–15/DR18–23/DC to the gastrocnemius muscle of 2-month-old mdx mice. The sarcolemmal integrity and central nucleation were examined at 40 and 60 days after gene transfer, respectively (transduction efficiency was ~80%). Expressing the DR2–15/DR18–23/DC microgene efficiently prevented sarcolemmal leakage, as evidenced by the lack of Evans blue dye accumulation in transduced myofibers (Figure 3A). The percentage of central nucleation reflects pathogenic muscle degeneration/regeneration. We observed a significant reduction in central nucleation in the DR2–15/DR18–23/DC microgene–positive myofibers (Figure 3A and Supplemental Figure 5).

Figure 3

The R16/17-containing microgene ameliorates muscle disease and improves muscle force following AAV gene transfer. (A) Results from adult mdx muscle (local delivery; n = 4–8 for each assay). Top: Serial sections with H&E and nNOS activity staining (see immunostaining in Supplemental Figure 5); scale bar: 100 μm. Bottom left: Evans blue dye (EBD) uptake; scale bar: 400 μm. An R16-specific antibody revealed the microgene. Bottom right: Central nuclear quantification. (B) Specific tetanic force in the mdx EDL muscle following systemic neonatal microgene therapy. ^#P ≤ 0.033 compared with all other groups; ^†P ≤ 0.001 compared with mdx only. (C) Systemic delivery of AV.CMV.DR2–15/DR18–23/DC to adult MyoD/dystrophin double-knockout (M-dko) mice improved skeletal muscle function and reduced the serum CK level. Sample size for muscle function assay: n = 4 for uninfected littermate controls; n = 14 for AV.CMV.DR2–15/DR18–23/DC–infected mice. Sample size for the CK assay: n = 3 for uninfected littermate controls; n = 6 for AV.CMV.DR2–15/DR18–23/DC–infected mice. *P ≤ 0.002, ^‡P = 0.015 compared with controls.

Next, we examined muscle strength. We have previously established the DR4–23/DC microgene as the best candidate microgene (16, 26, 27). Here we compared muscle force in DR4–23/DC and DR2–15/DR18–23/DC microgene–treated mdx mice. We delivered AAV-9 AV.CMV.DR4–23/DC or AV.CMV.DR2–15/DR18–23/DC to newborn mdx mice through facial vein injection. At 3 months after systemic gene transfer, we examined microgene expression and muscle force in the extensor digitorum longus (EDL) muscle (Figure 3B). Consistent with our previous reports (28), we observed highly efficient gene transfer with both AAV vectors (~80%, data not shown). Muscle force was also significantly enhanced by both microgenes (Figure 3B). Importantly, the extent of force improvement was identical.

To further evaluate the therapeutic effect of the DR2–15/DR18–23/DC microgene, we performed systemic AAV delivery in mdx4cv mice. This model carries a different mutation in the dystrophin gene, and it is on a different genetic background (C57BL/6 rather than C57BL/10 [BL10]). Consistent with our findings in mdx mice, AV.CMV.DR2–15/DR18–23/DC significantly increased limb muscle strength and reduced serum creatine kinase (CK) levels in mdx4cv mice (Supplemental Figure 6).

In contrast to those in human patients, the clinical manifestations are quite mild in both mdx and mdx4cv mice. This is partially due to robust muscle regeneration. MyoD is a master transcription regulator for skeletal muscle differentiation. Eliminating MyoD abolishes muscle regeneration in mdx mice. As a result, MyoD-null mdx mice develop a more severe skeletal muscle disease that better models the human condition (29). To validate the therapeutic efficacy of the DR2–15/DR18–23/DC microgene in a symptomatic model, we delivered AAV-9 AV.CMV.DR2–15/DR18–23/DC to 2-month-old MyoD-null mdx mice through tail vein injection. Three months after systemic gene transfer, the EDL muscle specific force was significantly enhanced in AAV-treated mice (Figure 3C). Furthermore, eccentric contraction–induced muscle injury was mitigated, and serum CK levels were significantly reduced (Figure 3C). Taken together, the results indicate that the R16/17-containing microgene AAV vector recovered sarcolemmal nNOS and ameliorated muscle pathology.

The R16/17-containing minigene improves vascular perfusion in contracting muscle and enhances exercise performance. The minigenes are 60%–80% larger than the microgenes, and they yield better functional recovery in mdx mice (25, 26). However, none of the current minigenes contain R16/17, nor can they restore nNOS. Here, we tested whether adding R16/17 could further improve the therapeutic efficacy of the minigene. We generated DH2–R15 and DH2–R19 minigene transgenic mdx and mdx4cv mice (Figures 4, 5, 6, 7 and Supplemental Figures 7, 8, and 10). The DH2–R15 minigene carries R16/17, but the DH2–R19 minigene does not (Figure 1 and Table 1).

Figure 4

The R16/17-containing minigene (DH2–R15) restores sarcolemmal nNOS, normalizes muscle force to wild-type levels, and reduces eccentric contraction–associated muscle injury. (A) Representative Western blots of whole muscle lysates from the TA muscles of BL10, mdx, DH2–R15 transgenic mdx, DH2–R19 transgenic mdx, and nNOS-knockout mice. GAPDH was used as the loading control. (B) Representative Western blots of microsomal preparations from the quadriceps femoris muscles. (C) Representative photomicrographs of dystrophin immunostaining (H1, R16, R19) and nNOS activity staining from DH2–R15 and DH2–R19 transgenic mdx mice. Scale bar: 100 μm. (D) The DH2–R15 minigene normalizes muscle-specific force and prevents eccentric contraction-induced injury. *P ≤ 0.021 compared with transgenic mdx mice.

Figure 5

The DH2–R15 minigene attenuates norepinephrine-mediated vasoconstriction in contracting muscle. Decreases in femoral vascular conductance were recorded following intraarterial injection of 2 doses of norepinephrine (NE) into the resting and contracting hind limbs of male HSA(human skeletal α-actin promoter).DH2–R15 and HSA.DH2–R19 transgenic mdx mice. (A) Results from DH2–R15 transgenic mdx mice. (B) Results from DH2–R19 transgenic mdx mice. AUC is presented in arbitrary units. Asterisks indicate significant differences.

Figure 6

The DH2–R15 minigene transgenic mdx mice show better muscle perfusion during exercise. Muscle and internal organ perfusion at rest and during exercise was quantified using radiolabeled microspheres in male HSA.DH2–R15 and HSA.DH2–R19 transgenic mdx mice. (A) Tissue perfusion in selected muscles at rest. (B) Kidney perfusion at rest. (C) Tissue perfusion in selected muscles during exercise. (D) Kidney perfusion during exercise. Gastro., gastrocnemius muscle. Asterisks indicate significant differences.

Figure 7

The DH2–R15 minigene prevents ischemic injury and enhances performance during intensive exercise. (A) Body weight–normalized daily running distance in HSA.DH2–R15 and HSA.DH2–R19 transgenic mdx mice (6-month-old male mice). *P ≤ 0.02. (B and C) Representative photomicrographs of general histopathology (H&E staining), inflammation (macrophage infiltration), nNOS activity, and minidystrophin expression (immunostaining with epitope-specific antibodies) in the limb muscle harvested after 8 days’ intensive treadmill running. (B) Results from DH2–R15 transgenic mdx mice. Arrowhead indicates a single regenerating myofiber. (C) Results from DH2–R19 transgenic mdx mice. Arrows indicate macrophages. Scale bars: 50 μm. See Supplemental Figure 10 for low-magnification images.

On Western blot analysis, we observed minidystrophin bands of the expected size (Figure 4A). nNOS was detected in whole muscle lysate from both DH2–R19 and DH2–R15 transgenic mice (Figure 4A). However, in membrane-enriched microsomal preparation, it was detected only in DH2–R15 transgenic mice (Figure 4B). Immunofluorescence staining and in situ nNOS activity staining further confirmed Western blot results (Figure 4C). These studies revealed human minigene expression in both transgenic strains. The DH2–R15 transgenic mice also expressed R16, -17, -18, and -19, four repeats that were missing in DH2–R19 transgenic mice (Figure 4C; data not shown for R17 and R18 immunostaining). Importantly, nNOS expression and nNOS activity were only detected in muscle sections from DH2–R15 transgenic mice (Figure 4C; data not shown for nNOS immunostaining).

Muscle-specific force was completely restored to wild-type levels in both DH2–R19 and DH2–R15 minigene transgenic mice (Figure 4D) (26). When challenged with eccentric contraction, the 2 types of transgenic mice showed identical responses (Figure 4D). In the first 4 cycles, the force preservation was slightly lower than that in BL10 mice. From the fifth to the tenth cycle, there were no significant differences among the 3 groups (Figure 4D).

A critical function of membrane-associated nNOS is to generate NO to counteract α-adrenergic vasoconstriction during muscle contraction. The loss of this protective mechanism leads to functional ischemia and thus aggravates contraction-induced injury in dystrophin-deficient muscles (2, 3, 30). To determine whether the restoration of sarcolemmal nNOS by the R16/17-containing minigene could improve blood perfusion during muscle contraction, we compared femoral vascular conductance in DH2–R15 and DH2–R19 transgenic mdx and mdx4cv mice (Figure 5 and Supplemental Figures 7 and 8) (2). In resting muscle, the 2 strains displayed similar femoral vascular conductance (data not shown). When resting muscles were challenged with norepinephrine, the strains also showed similar reductions in femoral artery conductance (Figure 5 and Supplemental Figure 8). However, we observed a striking difference in contracting muscle. In DH2–R19 transgenic mice and transgene-negative littermates from DH2–R15 breeding, norepinephrine-induced vasoconstriction was not attenuated in the contracting muscles (Figure 5 and Supplemental Figure 7). In contrast, in DH2–R15 transgenic mice, the constrictor response provoked by norepinephrine administration was significantly reduced in the contracting muscles (Figure 5 and Supplemental Figure 8).

To further explore muscle perfusion in the intact animal, we directly measured blood flow using radiolabeled microspheres. We observed similar levels of kidney perfusion in DH2–R15 and DH2–R19 transgenic mice, irrespective of the activity status (resting or exercising) (Figure 6). The two strains also showed similar muscle perfusion at rest (Figure 6A). However, DH2–R15 transgenic mice demonstrated significantly greater muscle perfusion during the treadmill exercise (Figure 6B).

To measure the physiological benefit of nNOS restoration, we developed a treadmill protocol. Since muscle perfusion in sarcolemmal nNOS-deficient mice was compromised (Figure 5, Figure 6, and Supplemental Figures 7 and 8), we hypothesized that intensive daily exercise would lead to muscle ischemia in these mice. We further hypothesized that the accumulation of ischemic damage would reduce running performance at later time points. As expected, BL10 mice showed a gradual increase in running distance up to 10 days with daily exercise (the duration of the study), while the running distance of mdx mice leveled off after day 4 (Supplemental Figure 9). Although this finding supports our hypothesis, we cannot rule out the possibility that the inherent muscle pathology might have limited treadmill performance in mdx mice.

To exclude the potential influence of muscle disease, we next compared running performance in DH2–R15 and DH2–R19 transgenic mdx mice under a similar treadmill exercise regimen. We have shown that both DH2–R15 and DH2–R19 minigenes can efficiently ameliorate muscle pathology and improve muscle contraction (Figure 3A and Figure 4D) (25, 26). In this case, should we observe distinctive performance profiles, it would be more likely due to the difference in sarcolemmal nNOS localization between the 2 strains. Consistent with our hypothesis, treadmill performance was gradually improved in DH2–R15, but not in DH2–R19, transgenic mdx mice (Figure 7A). Starting from day 6, DH2–R19 transgenic mdx mice showed significantly shorter running distances than DH2–R15 transgenic mdx mice (Figure 7A). Our observation in DH2–R19 transgenic mdx mice was further confirmed in DR4–23 transgenic mdx mice (Supplemental Figure 9). Similar to the DH2–R19 minigene, the DR4–23 microgene cannot restore sarcolemmal nNOS. These mice showed a running profile similar to that of mdx mice (Supplemental Figure 9). Additional studies revealed less than 1% human dystrophin expression in the heart of all transgenic mdx mice (data not shown). Hence, the difference in exercise performance is unlikely due to accidental cardiac rescue.

To determine whether intensive daily exercise had indeed resulted in ischemic damage in DH2–R19 transgenic mouse muscle, we analyzed muscle histology at the end of the eighth day of exercise (Figure 7, B and C, and Supplemental Figure 10). Muscles from DH2–R15 transgenic mice appeared healthy, with occasionally centrally nucleated myofiber (<1%) (Figure 7B and Supplemental Figure 10). In contrast, we observed frequent focal myofiber degeneration and regeneration in muscles obtained from DH2–R19 transgenic mice (Figure 7C and Supplemental Figure 10). Importantly, we also detected prominent macrophage infiltration in these loci (Figure 7C). Taken together, our data strongly suggest that restoring sarcolemmal nNOS can improve muscle perfusion, prevent ischemic damage in contracting muscle, and enhance exercise performance.

nNOS plays a pivotal regulatory role in skeletal muscle physiology and disease (1). Interestingly, nNOS function depends on its location at the sarcolemma. Loss of sarcolemmal nNOS has been shown to induce functional ischemia and exaggerate exercise-associated fatigue (2, 3, 9, 10). For these reasons, there has been a great interest in understanding the mechanisms involved in anchoring nNOS to the sarcolemma.

Studies in the mid-1990s suggested that the correct positioning of nNOS requires the DGC, a multimeric protein complex consisting of dystrophin, dystroglycans, sarcoglycans, sarcospan, syntrophin, and dystrobrevin. Subsequent biochemical, structural, and genetic analyses demonstrated an essential role for syntrophin (7, 17–20). In particular, the PDZ domain of syntrophin forms a complex with the PDZ domain of nNOS (17, 18). At the same time, it was suggested that nNOS recruitment might require dystrophin. Since the dystrophin C-terminal domain contains a syntrophin binding site, the involvement of dystrophin seems quite logical. Surprisingly, sarcolemmal nNOS expression is not altered in mdx mice that express C-terminal domain–truncated dystrophins (15, 16). Furthermore, even in the presence of an intact C-terminal domain, nNOS is not detected at the sarcolemma (21, 25, 31). Taken together, these findings indicate that the dystrophin C-terminal domain may, at most, play a nominal role in nNOS recruitment.

A more puzzling issue is that nNOS is not always present on the sarcolemma even when syntrophin is there (16, 21, 23, 25). Apparently, additional help is needed to anchor nNOS to the sarcolemma. Several candidates have been proposed. These include other proteins in the DGC or other proteins that interact with the DGC, such as aquaporin-4, sodium channel, calcium pump, and caveolin-3 (32–34). Unfortunately, none of these have been confirmed.

Different regions in the dystrophin N-terminal and rod domains (such as exons 3–7 or 8, 10–42, 45, 45–47, and 51–52) have also been suggested to help recruit nNOS to the sarcolemma (21, 22, 35). Nevertheless, defining the nNOS recruiting domain in dystrophin has proven to be a quite challenging task. Several lines of evidence suggest that dystrophin may not bind to nNOS directly. First, it has been shown that nNOS interacts with other proteins mainly through the PDZ–PDZ domain interaction (36, 37). However, dystrophin does not contain a classic PDZ domain. Second, 2 independent studies have failed to detect a direct interaction between full-length dystrophin and nNOS (8, 21). Proving an indirect interaction is technically much more difficult. Third, it is not clear whether the dystrophin-nNOS interaction (assuming there is one) depends on a single defined region or several discontinuous regions in dystrophin.

In this study, we took a systemic approach to map the nNOS localization domain in dystrophin. We started with the DH2–R19 minigene (26). We have previously confirmed that this minigene cannot restore sarcolemmal nNOS (23, 25). Based on this information, we hypothesized that the H2-to-R19 region should contain the domain(s) required for nNOS recruiting. By in vivo plasmid transfection and AAV-mediated gene transfer, we narrowed down the nNOS recruiting domain to R16/17 (Figures 1 and 2, Supplemental Figures 2–4, and Table 1). We found that these 2 repeats were necessary to anchor nNOS to the sarcolemma. Furthermore, R16/17-mediated nNOS recruitment was not dependent on the flanking repeats or the dystrophin C-terminal domain. Interestingly, we also observed an interaction between the nNOS PDZ domain and R16/17, although its strength was lower than that of the interaction between the nNOS PDZ domain and the syntrophin PDZ domain (Figure 2, D and E). Based on these results, we here propose a novel nNOS recruitment model (Figure 8). According to this model, nNOS localization requires 2 independent interactions, one between R16/17 and the nNOS PDZ domain and the other between the syntrophin PDZ domain and the nNOS PDZ domain.

Figure 8

Schematic outline of sarcolemmal nNOS recruitment by an R16/17-containing microdystrophin gene. (A) The N-terminal domain (N) of the DR2–15/DR18–23/DC microgene interacts with γ-actin. The cysteine-rich domain (CR) of this microgene interacts with the dystroglycan complex (DG). Syntrophin (Syn) is brought to the sarcolemma by dystrobrevin (Dbr), which binds to the sarcoglycan complex (SG). nNOS recruitment requires 2 independent interactions, one between the nNOS PDZ β-finger and the syntrophin PDZ domain and the other between the nNOS PDZ groove and R16/17. Numbers denote spectrin-like repeats; H1 and H4, hinge 1 and hinge 4. (B) The hypothetical nNOS-R16/17 interaction model. The C-terminal helix of R16 joins the N-terminal helix of R17 to form a long helix. The R16/17 dimer then binds to the groove in the nNOS PDZ domain. Yellow, R16; blue, R17; dotted, flanking repeats.

This model raises several interesting questions. First, how can a single nNOS PDZ domain interact with 2 partners at the same time? A ternary complex involving the nNOS PDZ domain has been demonstrated before (34). However, in this type of the complex, the nNOS PDZ domain only interacts with one partner (such as CAPON and postsynaptic density proteins). This intermediate partner then recruits the third partner to the complex (such as small G protein Dexras 1, synapsins, and NMDA receptor) (34). The classical PDZ domain consists of 2 α helices (αA and αB) and 6 β-sheets (from βA to βF). αB and βB form a groove for ligand binding (37). The nNOS PDZ domain includes the first 150 N-terminal residues of nNOS. Amino acids 1–100 express the classic groove structure for ligand binding. The next 30 residues form a unique β-finger (17, 18). Interestingly, the syntrophin PDZ domain only binds to the β-finger. This leaves the nNOS PDZ groove unoccupied (17, 18, 38). Thus, there is a structural basis for the nNOS PDZ domain to bind to both syntrophin and R16/17 simultaneously (38).

Second, R16/17 does not seem to be a good ligand for the nNOS PDZ groove. The short C-terminal peptide motif and the internal peptide motif are the classical ligands for the PDZ groove (37). Dystrophin does not carry these motifs. It is currently unclear how spectrin-like repeats may interact with the PDZ domain. Interestingly, such interaction has also been observed in another PDZ domain–containing protein. Xia et al. have demonstrated that the PDZ domain of the actinin-associated LIM protein binds to spectrin-like repeats in α-actinin-2 (39). Interestingly, this interaction also requires 2 complete repeats (39). Taking together, these findings reveal a new recognition mode for the PDZ domain. The PDZ domain is one of the most frequently used scaffolding modules. Approximately 400 PDZ domains have been identified in the human genome. The identification/confirmation of spectrin-like repeats as a PDZ domain ligand sheds new light on our understanding of protein-protein interaction and oligomerization.

R16/17 is encoded by exons 41–46 (40). This explains the findings seen in the majority of the clinical cases. In these patients, the loss of sarcolemmal nNOS correlates with mutations between exons 41 and 46 (21, 22, 35). However, sarcolemmal nNOS is also missing in some patients who were reported to carry mutations in other regions of the dystrophin gene, such as exons 3–8 and 48–52 (21, 22, 35). There are at least 2 possibilities. Spectrin-like repeats are modular repeating elements. Each repeat is composed of 3 α-helices linked by turns (41). However, the structural unit of the dystrophin rod domain is not based on individual repeats. Instead, it is believed that the biophysical unit is formed by adjacent repeats in a nested manner (Figure 8B) (42). Disruption of one repeat may alter structural conformation of the nearby units and influence the correct phasing of the entire rod domain (43). Hence, mutations in other exons may alter the noncanonical PDZ groove binding motif formed by R16/17. Alternatively, it is also possible that these patients carry additional small mutations (such as single nucleotide change) between exons 41 and 46.

The ultimate goal of our study is to develop an effective gene therapy to treat all affected muscles in DMD/BMD. AAV is the only viral vector proven to be capable of body-wide gene delivery (27, 44). However, AAV cannot carry the full-length dystrophin coding sequence. Minimizing the dystrophin gene offers a valid avenue to circumvent this problem (26). A variety of different minigenes and microgenes have been developed. Unfortunately, none of the abbreviated genes have been conclusively shown to recruit nNOS to the sarcolemma.

Based on the finding that R16/17 is sufficient to restore sarcolemmal nNOS, we generated a new microdystrophin gene, DR2–15/DR18–23/DC. This microgene is similar to our previously reported DR4–23/DC microgene, except that R2–H2 is replaced by R16/17 (Figure 2A) (16, 26). Local and systemic gene transfer studies were performed in newborn and adult mouse DMD models. In mdx, mdx4cv, and MyoD/dystrophin double-knockout mice, the R16/17-containing microgene not only restored sarcolemmal nNOS, but also effectively reduced muscle histopathology and enhanced sarcolemma integrity (Figure 2, Figure 3, and Supplemental Figures 4 and 5). Furthermore, muscle strength and resistance to contraction-induced injury were improved to levels comparable to those resulting from DR4–23/DC microgene treatment (Figure 3B and Supplemental Figure 6).

One caveat with the microgenes is the uncertainty of their function in the canine animal model and human patients (45). On the other hand, the minigenes are derived from a remarkably functional template isolated from a patient with very mild BMD (46). They also restore muscle-specific forces to normal levels (Figure 4D) (26). Importantly, the minigenes can be delivered by trans-splicing AAV vectors to all muscles in the body (25, 28). From this standpoint, minigene therapy could be more effective than microgene therapy. To determine the therapeutic advantage of the R16/17-containing minigene, we next compared muscle force, femoral artery hemodynamics, muscle perfusion, and treadmill running in DH2–R15 and DH2–R19 transgenic mice (Figures 4–7 and Supplemental Figures 7, 8, and 10). Both transgenic strains were able to recover wild-type-level muscle force (Figure 4D). However, only the R16/17-containing minigene transgenic mice (DH2–R15) showed significantly improved perfusion in contracting muscle (Figure 5, Figure 6, and Supplemental Figure 8). Furthermore, restoring sarcolemmal nNOS by transfection of the DH2–R15 minigene significantly enhanced exercise performance and prevented ischemic damage (Figure 7 and Supplemental Figure 10).

Taken together, our studies provide evidence that R16/17 is essential for anchoring nNOS to the sarcolemma. The R16/17-containing mini-/microgenes may be more effective for DMD gene therapy than all previously described mini-/microgenes. Our findings also shed new light on the molecular mechanisms of nNOS PDZ domain–mediated scaffolding. Future studies on how R16/17 coordinates with syntrophin to bring nNOS to the sarcolemma may unravel yet-unrecognized aspects of this fascinating biological process.

View Supplemental data

We thank Glenn Morris, Lam Le, Stanley Froehner, Marvin Adams, David Bredt, Samie Jaffrey, and Michael Rudnicki for providing experiment reagents. We thank Elizabeth Critser and Natalia Karasseva for help with generating the founder transgenic mice. We thank John Cannon, Deyu Fang, and Harold Laughlin for helpful discussion. This work was supported by grants from the NIH (AR49419, AR57209, NS62934, and HL37387 to D. Duan, J.S. Chamberlain, G.D. Thomas, H.T. Yang, R.L. Terjung) and the Muscular Dystrophy Association (to D. Duan and J.S. Chamberlain).

Address correspondence to: Dongsheng Duan, Department of Molecular Microbiology and Immunology, University of Missouri, One Hospital Dr., Columbia, Missouri 65212, USA. Phone: (573) 884-9584; Fax: (573) 882-4287; E-mail: duand@missouri.edu.

Conflict of interest: The authors have declared that no conflict of interest exists.

Nonstandard abbreviations used: AAV, adeno-associated virus; BMD, Becker muscular dystrophy; BL10, C57BL/10; C, dystrophin C-terminal domain; CK, creatine kinase; DGC, dystrophin-associated glycoprotein complex; DMD, Duchenne muscular dystrophy; EDL, extensor digitorum longus; H, hinge region in the dystrophin rod domain; nNOS, neuronal NOS; PDZ, postsynaptic density 95, discs large, and zonula occludens–1; R, dystrophin spectrin-like repeat; TA, tibialis anterior.

Reference information: J. Clin. Invest.119:624–635 (2009). doi:10.1172/JCI36612

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1. Stamler, J.S., Meissner, G. 2001. Physiology of nitric oxide in skeletal muscle. Physiol. Rev. 81:209-237.
   View this article via: PubMed Google Scholar
2. Thomas, G.D., et al. 1998. Impaired metabolic modulation of alpha-adrenergic vasoconstriction in dystrophin-deficient skeletal muscle. Proc. Natl. Acad. Sci. U. S. A. 95:15090-15095.
   View this article via: CrossRef PubMed Google Scholar
3. Sander, M., et al. 2000. Functional muscle ischemia in neuronal nitric oxide synthase-deficient skeletal muscle of children with Duchenne muscular dystrophy. Proc. Natl. Acad. Sci. U. S. A. 97:13818-13823.
   View this article via: CrossRef PubMed Google Scholar
4. Kunkel, L.M. 2005. 2004 William Allan Award address. Cloning of the DMD gene. Am. J. Hum. Genet. 76:205-214.
   View this article via: CrossRef PubMed Google Scholar
5. Mendell, J.R., Engel, W.K., Derrer, E.C. 1971. Duchenne muscular dystrophy: functional ischemia reproduces its characteristic lesions. Science. 172:1143-1145.
   View this article via: CrossRef PubMed Google Scholar
6. Parker, J.M., Mendell, J.R. 1974. Proximal myopathy induced by 5-HT-imipramine simulates Duchenne dystrophy. Nature. 247:103-104.
   View this article via: PubMed Google Scholar
7. Brenman, J.E., Chao, D.S., Xia, H., Aldape, K., Bredt, D.S. 1995. Nitric oxide synthase complexed with dystrophin and absent from skeletal muscle sarcolemma in Duchenne muscular dystrophy. Cell. 82:743-752.
   View this article via: CrossRef PubMed Google Scholar
8. Chang, W.J., et al. 1996. Neuronal nitric oxide synthase and dystrophin-deficient muscular dystrophy. Proc. Natl. Acad. Sci. U. S. A. 93:9142-9147.
   View this article via: CrossRef PubMed Google Scholar
9. Percival, J.M., et al. 2008. Functional deficits in nNOSmu-deficient skeletal muscle: myopathy in nNOS knockout mice. PLoS ONE. 3:e3387.
   View this article via: PubMed Google Scholar
10. Kobayashi, Y.M., et al. 2008. Sarcolemma-localized nNOS is required to maintain activity after mild exercise. Nature. 456:511-515.
    View this article via: CrossRef PubMed Google Scholar
11. Ahn, A.H., Kunkel, L.M. 1995. Syntrophin binds to an alternatively spliced exon of dystrophin. J. Cell Biol. 128:363-371.
    View this article via: CrossRef PubMed Google Scholar
12. Suzuki, A., Yoshida, M., Ozawa, E. 1995. Mammalian alpha 1- and beta 1-syntrophin bind to the alternative splice- prone region of the dystrophin COOH terminus. J. Cell Biol. 128:373-381.
    View this article via: CrossRef PubMed Google Scholar
13. Yang, B., Jung, D., Rafael, J.A., Chamberlain, J.S., Campbell, K.P. 1995. Identification of alpha-syntrophin binding to syntrophin triplet, dystrophin, and utrophin. J. Biol. Chem. 270:4975-4978.
    View this article via: CrossRef PubMed Google Scholar
14. Yoshida, M., et al. 2000. Biochemical evidence for association of dystrobrevin with the sarcoglycan-sarcospan complex as a basis for understanding sarcoglycanopathy. Hum. Mol. Genet. 9:1033-1040.
    View this article via: CrossRef PubMed Google Scholar
15. Crawford, G.E., et al. 2000. Assembly of the dystrophin-associated protein complex does not require the dystrophin COOH-terminal domain. J. Cell Biol. 150:1399-1410.
    View this article via: CrossRef PubMed Google Scholar
16. Yue, Y., Liu, M., Duan, D. 2006. C-terminal truncated microdystrophin recruits dystrobrevin and syntrophin to the dystrophin-associated glycoprotein complex and reduces muscular dystrophy in symptomatic utrophin/dystrophin double knock-out mice. Mol. Ther. 14:79-87.
    View this article via: CrossRef PubMed Google Scholar
17. Hillier, B.J., et al. 1999. Unexpected modes of PDZ domain scaffolding revealed by structure of nNOS-syntrophin complex. Science. 284:812-815.
    View this article via: CrossRef PubMed Google Scholar
18. Tochio, H., Zhang, Q., Mandal, P., Li, M., Zhang, M. 1999. Solution structure of the extended neuronal nitric oxide synthase PDZ domain complexed with an associated peptide. Nat. Struct. Biol. 6:417-421.
    View this article via: CrossRef PubMed Google Scholar
19. Kameya, S., et al. 1999. alpha1-syntrophin gene disruption results in the absence of neuronal- type nitric-oxide synthase at the sarcolemma but does not induce muscle degeneration. J. Biol. Chem. 274:2193-2200.
    View this article via: CrossRef PubMed Google Scholar
20. Adams, M.E., Mueller, H.A., Froehner, S.C. 2001. In vivo requirement of the alpha-syntrophin PDZ domain for the sarcolemmal localization of nNOS and aquaporin-4. J. Cell Biol. 155:113-122.
    View this article via: CrossRef PubMed Google Scholar
21. Chao, D.S., et al. 1996. Selective loss of sarcolemmal nitric oxide synthase in Becker muscular dystrophy. J. Exp. Med. 184:609-618.
    View this article via: CrossRef PubMed Google Scholar
22. Wells, K.E., et al. 2003. Relocalization of neuronal nitric oxide synthase (nNOS) as a marker for complete restoration of the dystrophin associated protein complex in skeletal muscle. Neuromuscul. Disord. 13:21-31.
    View this article via: CrossRef PubMed Google Scholar
23. Judge, L.M., Haraguchi, M., Chamberlain, J.S. 2006. Dissecting the signaling and mechanical functions of the dystrophin-glycoprotein complex. J. Cell Sci. 119:1537-1546.
    View this article via: CrossRef PubMed Google Scholar
24. Grady, R.M., et al. 1999. Role for alpha-dystrobrevin in the pathogenesis of dystrophin-dependent muscular dystrophies. Nat. Cell Biol. 1:215-220.
    View this article via: CrossRef PubMed Google Scholar
25. Lai, Y., et al. 2005. Efficient in vivo gene expression by trans-splicing adeno-associated viral vectors. Nat. Biotechnol. 23:1435-1439.
    View this article via: CrossRef PubMed Google Scholar
26. Harper, S.Q., et al. 2002. Modular flexibility of dystrophin: implications for gene therapy of Duchenne muscular dystrophy. Nat. Med. 8:253-261.
    View this article via: CrossRef PubMed Google Scholar
27. Gregorevic, P., et al. 2004. Systemic delivery of genes to striated muscles using adeno-associated viral vectors. Nat. Med. 10:828-834.
    View this article via: CrossRef PubMed Google Scholar
28. Ghosh, A., Yue, Y., Long, C., Bostick, B., Duan, D. 2007. Efficient whole-body transduction with trans-splicing adeno-associated viral vectors. Mol. Ther. 15:750-755.
    View this article via: PubMed Google Scholar
29. Megeney, L.A., Kablar, B., Garrett, K., Anderson, J.E., Rudnicki, M.A. 1996. MyoD is required for myogenic stem cell function in adult skeletal muscle. Genes Dev. 10:1173-1183.
    View this article via: CrossRef PubMed Google Scholar
30. Dudley, R.W., et al. 2006. Sarcolemmal damage in dystrophin deficiency is modulated by synergistic interactions between mechanical and oxidative/nitrosative stresses. Am. J. Pathol. 168:1276-1287.
    View this article via: CrossRef PubMed Google Scholar
31. Li, S., et al. 2006. A highly functional mini-dystrophin/GFP fusion gene for cell and gene therapy studies of Duchenne muscular dystrophy. Hum. Mol. Genet. 15:1610-1622.
    View this article via: CrossRef PubMed Google Scholar
32. Miyagoe-Suzuki, Y., Takeda, S.I. 2001. Association of neuronal nitric oxide synthase (nNOS) with alpha1-syntrophin at the sarcolemma. Microsc. Res. Tech. 55:164-170.
    View this article via: CrossRef PubMed Google Scholar
33. Albrecht, D.E., Froehner, S.C. 2002. Syntrophins and dystrobrevins: defining the dystrophin scaffold at synapses. Neurosignals. 11:123-129.
    View this article via: CrossRef PubMed Google Scholar
34. Kone, B.C., Kuncewicz, T., Zhang, W., Yu, Z.Y. 2003. Protein interactions with nitric oxide synthases: controlling the right time, the right place, and the right amount of nitric oxide. Am. J. Physiol. Renal Physiol. 285:F178-F190.
    View this article via: PubMed Google Scholar
35. Torelli, S., et al. 2004. Absence of neuronal nitric oxide synthase (nNOS) as a pathological marker for the diagnosis of Becker muscular dystrophy with rod domain deletions. Neuropathol. Appl. Neurobiol. 30:540-545.
    View this article via: CrossRef PubMed Google Scholar
36. Kone, B.C. 2000. Protein-protein interactions controlling nitric oxide synthases. Acta Physiol. Scand. 168:27-31.
    View this article via: CrossRef PubMed Google Scholar
37. Harris, B.Z., Lim, W.A. 2001. Mechanism and role of PDZ domains in signaling complex assembly. J. Cell. Sci. 114:3219-3231.
    View this article via: PubMed Google Scholar
38. Abdelmoity, A., Padre, R.C., Burzynski, K.E., Stull, J.T., Lau, K.S. 2000. Neuronal nitric oxide synthase localizes through multiple structural motifs to the sarcolemma in mouse myotubes. FEBS Lett. 482:65-70.
    View this article via: CrossRef PubMed Google Scholar
39. Xia, H., Winokur, S.T., Kuo, W.L., Altherr, M.R., Bredt, D.S. 1997. Actinin-associated LIM protein: identification of a domain interaction between PDZ and spectrin-like repeat motifs. J. Cell Biol. 139:507-515.
    View this article via: CrossRef PubMed Google Scholar
40. Roberts, R.G., Coffey, A.J., Bobrow, M., Bentley, D.R. 1993. Exon structure of the human dystrophin gene. Genomics. 16:536-538.
    View this article via: CrossRef PubMed Google Scholar
41. Djinovic-Carugo, K., Young, P., Gautel, M., Saraste, M. 1999. Structure of the alpha-actinin rod: molecular basis for cross-linking of actin filaments. Cell. 98:537-546.
    View this article via: CrossRef PubMed Google Scholar
42. Calvert, R., Kahana, E., Gratzer, W.B. 1996. Stability of the dystrophin rod domain fold: evidence for nested repeating units. Biophys. J. 71:1605-1610.
    View this article via: CrossRef PubMed Google Scholar
43. Menhart, N. 2006. Hybrid spectrin type repeats produced by exon-skipping in dystrophin. Biochim. Biophys. Acta. 1764:993-999.
    View this article via: PubMed Google Scholar
44. Yue, Y., et al. 2008. A single intravenous injection of adeno-associated virus serotype-9 leads to whole body skeletal muscle transduction in dogs. Mol. Ther. 16:1944-1952.
    View this article via: CrossRef PubMed Google Scholar
45. Sampaolesi, M., et al. 2006. Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs. Nature. 444:574-579.
    View this article via: CrossRef PubMed Google Scholar
46. England, S.B., et al. 1990. Very mild muscular dystrophy associated with the deletion of 46% of dystrophin. Nature. 343:180-182.
    View this article via: CrossRef PubMed Google Scholar
47. Ausubel, F.M., et al., editors. 1995.Current protocols in molecular biology. John Wiley and Sons. New York, New York, USA.
48. Thanh, L.T., Nguyen, T.M., Helliwell, T.R., Morris, G.E. 1995. Characterization of revertant muscle fibers in Duchenne muscular dystrophy, using exon-specific monoclonal antibodies against dystrophin. Am. J. Hum. Genet. 56:725-731.
    View this article via: PubMed Google Scholar
49. Hope, B.T., Michael, G.J., Knigge, K.M., Vincent, S.R. 1991. Neuronal NADPH diaphorase is a nitric oxide synthase. Proc. Natl. Acad. Sci. U. S. A. 88:2811-2814.
    View this article via: CrossRef PubMed Google Scholar
50. Bulfield, G., Siller, W.G., Wight, P.A., Moore, K.J. 1984. X chromosome-linked muscular dystrophy (mdx) in the mouse. Proc. Natl. Acad. Sci. U. S. A. 81:1189-1192.
    View this article via: CrossRef PubMed Google Scholar
51. Chapman, V.M., Miller, D.R., Armstrong, D., Caskey, C.T. 1989. Recovery of induced mutations for X chromosome-linked muscular dystrophy in mice. Proc. Natl. Acad. Sci. U. S. A. 86:1292-1296.
    View this article via: CrossRef PubMed Google Scholar
52. Li, D., Yue, Y., Duan, D. 2008. Preservation of muscle force in mdx3cv mice correlates with low-level expression of a near full-length dystrophin protein. Am. J. Pathol. 172:1332-1341.
    View this article via: CrossRef PubMed Google Scholar
53. Ervasti, J.M., Campbell, K.P. 1991. Membrane organization of the dystrophin-glycoprotein complex. Cell. 66:1121-1131.
    View this article via: CrossRef PubMed Google Scholar
54. Yang, H.T., Dinn, R.F., Terjung, R.L. 1990. Training increases muscle blood flow in rats with peripheral arterial insufficiency. J. Appl. Physiol. 69:1353-1359.
    View this article via: PubMed Google Scholar
55. Flaim, S.F., Zelis, R.F. 1980. Regional distribution of cardiac output in conscious rats at rest and during exercise. Effects of diltiazem. Chest. 78:187-192.
    View this article via: PubMed Google Scholar
56. Janssen, B., Debets, J., Leenders, P., Smits, J. 2002. Chronic measurement of cardiac output in conscious mice. Am. J. Physiol. Regul. Integr. Comp. Physiol. 282:R928-R935.