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Human Molecular Genetics Advance Access originally published online on July 6, 2004
Human Molecular Genetics 2004 13(17):1873-1884; doi:10.1093/hmg/ddh204
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Human Molecular Genetics, Vol. 13, No. 17 © Oxford University Press 2004; all rights reserved

Defects in neuromuscular junction structure in dystrophic muscle are corrected by expression of a NOS transgene in dystrophin-deficient muscles, but not in muscles lacking {alpha}- and ß1-syntrophins

Terry Shiao1, Andrew Fond1, Bo Deng1, Michelle Wehling-Henricks1, Marvin E. Adams3, Stanley C. Froehner3 and James G. Tidball1,2,*

1Department of Physiological Science and 2Department of Pathology and Laboratory Medicine, University of California, Los Angeles, CA, USA and 3Department of Physiology and Biophysics, University of Washington, Seattle, WA, USA

Received April 1, 2004; Accepted June 21, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Muscular dystrophies that arise from mutations of genes that encode proteins in the dystrophin–glycoprotein complex (DGC) frequently involve defects in the structure of neuromuscular junctions (NMJs). DGC mutations that cause NMJ defects typically cause a secondary loss of neuronal nitric oxide synthase (nNOS) from the post-synaptic membrane. We tested the hypothesis that reduction of muscle-derived NO production causes NMJ defects in DGC mutants by analyzing the effect of modulating muscle NO production on NMJ structure in mutant and wild-type muscles. We found that nNOS null mutants, dystrophin-deficient mdx mice and {alpha}-syntrophin null mutants showed reductions in the concentration of acetylcholine receptors (AChRs) at the post-synaptic membrane. Also, expression of a muscle-specific NOS transgene increased AChR concentration, which reflected an increase in both AChR expression and clustering. NOS transgene expression also increased the size of NMJs, and partially corrected defects in normal NMJ architecture that were observed in mdx and {alpha}-syntrophin null muscles. In addition, stimulation of AChR clustering in vitro by application of laminin or VVA B4 lectin induced a 3–4-fold increase in NOS activity and increased AChR clustering that could be prevented by NOS inhibition. However, the partial rescue of NMJ structure by expression of a NOS transgene required the expression of {alpha}- or ß1-syntrophin at the NMJ; partial NMJ rescue was seen in the muscles of {alpha}-syntrophin mutants that expressed ß1-syntrophin, but no rescue was observed in muscles of {alpha}-syntrophin mutants that also lacked ß1-syntrophin. These findings show that NO promotes AChR expression and clustering in vivo and contributes to normal NMJ architecture. The results suggest that defects in NMJ structure that occur in some DGC mutants can result from the secondary loss of NOS from muscle.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Defects in the expression of members of the dystrophin–glycoprotein complex (DGC) typically disrupt the structure of the neuromuscular junction (NMJ). Loss of dystrophin has been most extensively examined in this regard, and shown to cause a great reduction in the folding of the post-synaptic membrane at NMJs, as well as leading to disruptions in the distribution of acetylcholine receptors (AChRs) and other proteins at the post-synaptic membrane (1–4). In addition, the concentration of functional AChRs at the post-synaptic membrane is reduced in dystrophin-deficient muscle, which is associated with impaired synaptic transmission (5). The architectural and functional deficiencies of dystrophin-deficient NMJs may reflect loss of structural contributions of the dystrophin complex to forming and maintaining normal NMJs. For example, a role for the dystrophin complex in mediating lateral associations between the actin cytoskeleton and folds in the cell membrane has been inferred from electron microscopic observations (6), suggesting that reduced post-synaptic folding and other structural defects may be consequences of reductions in those lateral associations.

Surprisingly, the loss of utrophin, a close homologue of dystrophin that is also concentrated at NMJs, produces only slight defects in the architecture of the post-synaptic membrane, and yields small reductions in the concentration or distribution of AChRs at the NMJ (7,8). Because utrophin null mutant mice display minor NMJ defects and negligible muscle pathology, whereas dystrophin-deficient mdx muscles show extensive pathology and NMJ defects, it is possible that mdx NMJ defects are a non-specific effect of muscle injury. The interpretation is further supported by the observation that NMJs of wild-type muscles that have regenerated following toxin injection show disruptions of NMJ structure (9). However, several findings show that disruptions of the NMJ in muscles with DGC defects cannot be explained as only the non-specific consequences of muscle repair following injury. For example, the expression of a truncated dystrophin in muscles of dystrophin-deficient and utrophin-deficient double-mutant mice reduced NMJ structural defects without reducing muscle pathology (4).

Null mutations for genes that encode other members of the DGC also cause defects in NMJ structure, but only minor muscle pathology. Loss of {alpha}-dystrobrevin from the dystrophin complex causes a reduction of folding of the post-synaptic membrane, although there is no apparent disruption in the distribution or concentration of either dystrophin or utrophin at the post-synaptic membrane (10). The muscles of {alpha}-dystrobrevin null mice also show abnormal distribution of AChRs at the post-synaptic membrane in vivo, and display defects in clustering of AChRs on myotubes that were stimulated in vitro with a lectin that binds N-acetylgalactosamine-terminated glycoconjugates (10). The finding that defects in AChR clustering in muscles lacking {alpha}-dystrobrevin resembles clustering defects in mdx muscles that lack the entire dystrophin complex has been interpreted as showing that {alpha}-dystrobrevin may play a key role in mediating the functions of the DGC at the post-synaptic membrane (10).

More recently, null mutation in the dystrophin-binding partner {alpha}-syntrophin has also been shown to cause pronounced reductions in folding of the post-synaptic membrane at NMJs (11,12), although the {alpha}-syntrophin null mutants do not show detectable muscle damage or regeneration (11). Interestingly, utrophin is also lost from the post-synaptic membrane of {alpha}-syntrophin null mice. Although the mechanisms through which {alpha}-syntrophin-deficiency leads to defects in NMJ structure and loss of utrophin from the NMJ are unknown, these findings have suggested that {alpha}-syntrophin may also be involved in signaling that is required for the normal assembly of NMJs.

However, other members of the DGC can be lost through null mutation without leading to any detectable defect in NMJ structure. Loss of either {alpha}-sarcoglycan or {gamma}-sarcoglycan produces no apparent defect in NMJ structure, although both sarcoglycan mutants undergo muscle damage and regeneration (13,14). Thus, disruptions of the architecture of NMJs are not a generic consequence of perturbations of DGC structure, and must result from loss of function associated with specific members of the DGC.

The DGC members whose loss leads to disruptions of NMJ structure share the common characteristic that they are ligands for neuronal nitric oxide synthase (nNOS), or they are responsible for localizing NOS ligands at the NMJ. NOS binds to {alpha}- or ß1-syntrophin through its PDZ domain (15–17). The syntrophins in turn are concentrated at the post-synaptic membrane by binding at the C-terminal regions of dystrophin, utrophin or dystrobrevin (18–20). Loss of expression of any of these DGC members that bind directly or indirectly to NOS, leads to a reduction of NOS from the muscle cell membrane (11,21–23). In addition, loss of DGC members that do not play a role in NOS localization at the membrane, produces no defects in NMJ structure (13,14). The coincidence of the loss of direct or indirect binding partners of nNOS from the muscle cell membrane and the occurrence of defects in NMJ structure support the hypothesis that defects in NOS localization or NO production may contribute to defects in NMJ structure.

In the present investigation, we test the hypothesis that nNOS plays a role in regulating the structure of NMJs in vivo. We also evaluate whether muscle-derived NO can promote the expression or clustering of AChR at mature or incipient NMJs. We test this hypothesis in several lines of genetically modified or spontaneously mutant mice that include: (1) mdx mice that lack dystrophin and have greatly reduced levels of expression of other DGC members; (2) nNOS null mutant mice; (3) {alpha}-syntrophin null mutant mice; (4) nNOS transgenic mice; (5) mdx mice that express a NOS transgene; and (6) {alpha}-syntrophin null mice that express a NOS transgene. Collectively, our findings indicate that muscle-derived NO increases the expression and clustering of AChRs, and increases both the size of NMJs and the concentration of AChRs at the post-synaptic membrane. Furthermore, we find that normalizing NO production in mdx mice by the expression of a NOS transgene greatly reduces the defects in NMJ structure, and that NOS is localized at the NMJs of these animals. Similarly, NOS transgene expression by {alpha}-syntrophin –/– muscle that expresses ß1-syntrophin causes an increase in NOS at the NMJ and produces a small but significant improvement in NMJ structure. However, expression of a NOS transgene in muscles that lack {alpha}- and ß1-syntrophin causes an accumulation of NOS in the cytosol rather than at the cell membrane, and does not improve NMJ structure. These observations support our conclusion that NOS plays a role in regulating NMJ structure, but that its role in regulating NMJ structure also requires syntrophin expression.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Expression of a NOS transgene increases expression of AChR
Western analysis of membrane fractions from muscles of nNOS Tg/mdx mice and non-Tg mdx mice confirmed that NOS transgene expression increased the concentration of nNOS at the cell membrane (Fig. 1). Western analysis also showed that the expression of the nNOS transgene increases the concentration of AChR-{alpha} in the muscle cell membrane by ~190% relative to non-Tg mdx mice. In addition, expression profiling data showed that there is a 17-fold increase in AChR-{alpha} mRNA in the muscles of NOS Tg mice compared with non-transgenic littermates, and a 170% increase in the concentration of AChR-{alpha} mRNA in the muscles of NOS Tg/mdx mice compared with the non-Tg mdx. NOS Tg muscles showed an >250-fold increase in nNOS mRNA compared with non-transgenic littermates, and as assessed by expression profiling (24).



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  		Figure 1. Western blots of muscle membrane fractions. Each of the three left lanes was from a different, non-transgenic, mdx mouse. Each of the three right lanes was from a different NOS Tg/mdx mouse. (A) Anti-nNOS blot of mdx and NOS Tg/mdx muscles. With longer exposures, nNOS is detectable in mdx membrane fractions, which shows that there is an enrichment of nNOS in the NOS Tg/mdx membrane preparation by more than 50-fold. (B) Anti-AChR blot of non-Tg/mdx and NOS Tg/mdx muscle membranes.

 
Increased expression of NOS increases NMJ size and AChR density
Observations of whole mounts of individual muscle fibers labeled with fluoroscein isothionate {alpha}-bungarotoxin (FITC–BTX) indicated that modification of nNOS expression can affect NMJ structure and AChR receptor density (Fig. 2). In general, we observed that increases in NOS expression corresponded with increases in AChR density at the post-synaptic membrane. Fluorescent intensities of NMJs of FITC–BTX labeled fibers from mdx muscles (Fig. 2B), {alpha}-syn –/– muscles (Fig. 2C and D) or nNOS –/– muscles (Fig. 2E) were markedly less than C57 NMJs (Fig. 2A). NOS transgene expression in C57 muscle increased FITC–BTX labeling intensity (Fig. 2F), and NOS Tg/mdx NMJs showed increased labeling with FITC–BTX compared with mdx (Fig. 2G). Finally, NMJs of 2-week-old C57 (Fig. 2K) and 2-week-old mdx (Fig. 2L) showed no difference in FITC–BTX labeling intensity; previous investigations have shown that there is no significant difference in nNOS concentration in C57 and mdx muscles at this age (22).



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  		Figure 2. NMJs of individual muscle fibers labeled with FITC–BTX. (A) C57 EDL muscle, (B) mdx EDL muscle, (C) {alpha}-syn –/– EDL muscle, (D) {alpha}-syn –/– STM muscle, (E) NOS –/– EDL muscle, (F) NOS Tg EDL muscle, (G) NOS Tg/mdx EDL muscle, (H) NOS Tg/{alpha}-syn –/– EDL muscle, (I) NOS Tg/{alpha}-syn –/– STM muscle, (J) C57 SOL muscle, (K) 2-week-old C57 EDL muscle and (L) 2-week-old mdx EDL muscle. Bar=8 µm.

 
We quantitatively tested our qualitative observations by measuring the intensity of BTX fluorescence per unit area of NMJ (as a measure of AChR density), the length of post-synaptic membrane crests (as a measure of NMJ structure) and the surface area of the fiber occupied by the NMJ (as a measure of AChR dispersal). We found a strong effect of changes in nNOS expression on the density of AChRs, in which there is a 45% reduction (P<0.05) in AChR density in nNOS –/– mice, compared with the wild-type controls, and a 33% (P<0.05) increase in AChR density in NOS Tg mice compared with the non-Tg littermates (Fig. 3). A smaller difference in AChR density was also observed between C57 wild-type controls and non-Tg littermate controls of the NOS Tg mice, which may reflect differences between background strains. Importantly, NOS Tg mice that received the NOS inhibitor N-nitro-L-arginine methyl ester (L-NAME) in their drinking water showed AChR densities that were 60% less than untreated NOS Tg mice (P<0.05), and did not differ from nNOS –/– mice.



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  		Figure 3. Density of AChR receptors measured in en face views of individual fibers. Asterisk indicates significantly different from C57 value at P<0.05. Hash sign indicates significantly different from non-transgenic littermates (P<0.05). All data were obtained on EDL fibers, unless noted otherwise, where ‘Sol’ indicates soleus muscle and ‘Stm’ indicates sternomastoid muscle. Bars indicate standard error.

 
Similarly, we found that the length of post-synaptic membrane crests at the NMJ is positively related to NOS expression (Fig. 4). NOS –/– NMJs showed a small but significant decrease in the lengths of the post-synaptic membrane crests (13%; P<0.05), whereas NOS Tg mice showed a 41% increase (P<0.05) in crest length compared with the non-Tg littermates. The increase in crest length in NOS Tg mice was partially reversed by treatments with L-NAME (11% reduction; P<0.05). In most cases, levels of NOS expression also showed a positive relationship to the surface area of the fiber that was occupied by the NMJ. Null mutation of nNOS yielded a 24% reduction (P<0.05) in NMJ area, whereas NOS Tg mice showed a significant 26% increase (P<0.05) in the NMJ area that was reversed by treatment with L-NAME (Fig. 5). The finding that the NMJ area for mdx mice was greater than for C57 mice is an exception to the positive relationship between NOS expression and NMJ size. This likely reflects the broader dispersal of post-synaptic membrane on the surface of the mdx fibers that results from fragmentation of the NMJ.



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  		Figure 4. Length of post-synaptic membrane crests measured in en face views of individual fibers. Asterisk indicates significantly different from C57 value at P<0.05. All data were obtained on EDL fibers, unless noted otherwise, where ‘Sol’ indicates soleus muscle and ‘Stm’ indicates sternomastoid muscle. Bars indicate standard error.

 


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  		Figure 5. Area occupied by post-synaptic membrane measured in en face views of individual fibers. Asterisk indicates significantly different from C57 value at P<0.05. Hash sign indicates significantly different from non-transgenic littermates (P<0.05). All data were obtained on EDL fibers, unless noted otherwise, where ‘Sol’ indicates soleus muscle and ‘Stm’ indicates sternomastoid muscle. Bars indicate standard error.

 
Differences in AChR concentration between muscle fiber types correspond to differences in NOS expression
Because the concentration of AChRs at the NMJ was so strongly affected by null mutation or over-expression of nNOS, we tested whether similar differences in AChR concentration were observed between the NMJs of fast [extensor digitorum longus (EDL)] and slow muscle [soleus (SOL)]. Previous investigations have shown that nNOS is expressed at much higher levels in the fast fibers than in the slow fibers, in rodents (25). We found that AChR density in SOL muscle fibers was 25% less (P<0.05) than in fast EDL fibers from the same wild-type animals (Fig. 3B). In addition, the length of post-synaptic membrane crests at the NMJ of SOL fibers from wild-type animals was 31% less (P<0.05) than in EDL fibers (Fig. 4B). This suggests that some of the differences in NMJ architecture between fast and slow fibers may result from differences in the levels of nNOS expression.

Expression of a NOS transgene normalizes NMJ structure in mdx muscle
We tested whether the normalization of NO production in mdx muscle could repair defects in NMJ architecture and AChR concentration in mdx muscle. Our qualitative observations showed that expression of the NOS Tg in mdx muscle restored the normal, continuous-guttering of the post-synaptic membrane (Fig. 2G). In addition, measurements of AChR density at NOS Tg/mdx fiber membranes showed a 29% increase (P<0.05) in density compared with non-Tg mdx mice, although Tg expression did not recover AChR concentrations to wild-type levels (Fig. 3B). The lack of full return to wild-type levels of AChR concentration at the NMJs of NOS Tg/mdx fibers may result from other NMJ defects that are not related to NOS deficiency, or result from a failure to achieve wild-type levels of NO production at the post-synaptic membrane. Our findings also show that the expression of the NOS transgene in mdx mice has no significant effect on the length of post-synaptic membrane crests (Fig. 4B), although the surface area of the fiber that is occupied by the NMJ is increased significantly by the expression of the NOS transgene in the mdx muscle (29%; P<0.05) (Fig. 5B). Interestingly, the discontinuous-guttering of the post-synaptic membrane of adult mdx mice was not observed in pre-necrotic, 2-week-old mdx muscle fibers (Fig. 2L).

Expression of a NOS transgene partially rescues AChR density and post-synaptic membrane structure at the NMJs of {alpha}-syntrophin –/– muscles, only if ß1-syntrophin is present
Previous investigations (11) have shown that the muscles of {alpha}-syn –/– mice typically express ß1-syntrophin at the post-synaptic membrane, although sternomastoid (STM) muscles are unusual because there is an absence of both {alpha}- and ß1-syntrophin from the NMJs of {alpha}-syn –/– mice (26). We tested whether defects in NMJ structure and AChR clustering would result from expression of the nNOS transgene in {alpha}-syn –/– muscles that expressed or lacked ß1-syntrophin at the post-synaptic membrane. Our measurements show that {alpha}-syn –/– muscles show large reductions in AChR concentrations at the post-synaptic membranes of EDL and STM muscles (59 and 66% reductions, respectively) (Fig. 3C). Expression of the nNOS transgene in {alpha}-syn –/– EDL muscles (Fig. 2H) resulted in a significant, 45% increase in AChR concentration in EDL, but no change occurred in the STM (Figs 2I and 3C). Similarly, nNOS transgene expression in {alpha}-syn –/– muscles caused a small, but significant increase in the length of post-synaptic membrane crests of EDL, but not of STM muscles (Fig. 4C).

The inability of nNOS transgene expression to recover partially the structure of NMJs in {alpha}-syn –/– STM suggested the possibility that nNOS localization at the post-synaptic membrane may be required for modulating synapse structure. We observed that NOS Tg/mdx muscle contained elevated concentrations of nNOS at the NMJ, which was consistent with a relationship between localization and rescue. However, as previously reported, nNOS was not detectable at the post-synaptic membranes of either STM, quadriceps or EDL muscles in {alpha}-syn –/– mice (11). However, expression of the NOS transgene in {alpha}-syn –/– mice resulted in expression of nNOS at the NMJ and at extrasynaptic regions of the cell membrane of quadriceps and EDL, but not STM muscles (Fig. 6). Instead, NOS was distributed in a reticular pattern within the cytoplasm of NOS Tg/syn –/– fibers in the STM (Fig. 6D).



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  		Figure 6. Transverse sections of muscle from an {alpha}-syn –/– mouse expressing the NOS transgene. (A and B) Double-labeled section of a quadriceps muscle showing distribution of labeling with FITC–BTX (green) or anti-nNOS (red). (C and D) Double-labeled section of a STM muscle showing distribution of labeling with FITC–BTX (green) or anti-nNOS (red). Arrowheads indicate location of NMJ. Bar=25 µm.

 
Exogenous NO promotes AChR expression on the surface of myotubes in vitro
Application of any of the NO donors sodium nitroprusside (SNP), S-nitroso-N-acetylpenicilliamine (SNAP) or 2,2'-(hydroxynitrohydrazino)bis-ethanamine (NOC-18) produced significant and dose-dependent increases in the surface concentrations of AChRs on myotubes in vitro (Fig. 7A). However, application of the NOS inhibitor L-NAME to myotubes in vitro did not affect the increase in surface AChR concentration that occurs during normal, in vitro maturation of myotubes (Fig. 7B). This observation indicates that NO can promote surface expression of AChRs during myotube growth, but is not required.



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  		Figure 7. Measurement of fluorescent signal intensity of C2C12 myotubes after incubation with FITC–BTX. (A) Dose response of FITC-signal using three separate NO donors. Asterisk indicates significantly different from untreated, control myotubes. Bars indicate standard errors. (B) Timecourse of FITC-signal intensity for myotubes treated with 1.0 mM SNAP, or SNAP and L-NAME or untreated controls (DMEM). Asterisk indicates significantly different from untreated, control myotubes. Bars indicate standard errors.

 
Laminin or VVA B4 binding activates NOS and promotes AChR clustering in vitro
AChR clustering that is stimulated by laminin or Vicia villosa agglutinin B4 (VVA B4) on myotubes in vitro was prevented by NOS inhibition by L-NAME (Fig. 8A). In addition, we find that stimulation of myotubes with laminin or VVA B4 causes a large, significant increase in NO release by myotubes (Fig. 8B), which indicates that the binding of ligands that induce receptor clustering can indirectly increase NOS activation and thereby promote AChR clustering.



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  		Figure 8. Assays of AChR clustering and NO production by C2C12 myotubes with or without treatment with laminin or VVA B4. (A) Concentration of AChR on the surface of myotubes. (B) NO release from C2C12 myotubes. Asterisk indicates significantly different from untreated, control myotubes. Bars indicate standard errors.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Defects in the expression of any one of several members of the DGC have been shown to disrupt the organization of the post-synaptic membrane of NMJs, and to perturb AChR clustering (1–3,7,8,11,12,23,27,28). These observations have been interpreted as evidence that each of those DGC members plays a role in regulating the structure of NMJs, or participates in AChR clustering. However, nNOS at the NMJ is decreased as a consequence of the loss of many of these DGC proteins (10,11,21,22), so that an alternative and more parsimonius explanation for these NMJ defects would be that they arise from NOS deficiency. The major finding of the present investigation is that muscle-derived NO can modulate the expression and clustering of AChRs in vivo, and that defects in NMJ structure in dystrophin-deficient or {alpha}-syntrophin-deficient muscle can be reduced by expression of a muscle-specific NOS transgene. Thus, secondary defects in nNOS expression can contribute to disruptions in NMJ structure and function in dystrophinopathies and other pathologies that involve DGC defects.

The fragmented appearance of the NMJ of adult mdx mice is the most obvious defect in NMJ structure that is prevented by expression of the NOS transgene. Early investigators of NMJ structure in mdx muscle (1,2) noted that fragmentation of the NMJ became more severe with age, suggesting that the disruptions were secondary to necrosis and regeneration, rather than a direct consequence of dystrophin deficiency. Later findings showed that acute muscle injuries could produce fragmented NMJ appearance, which further supported the view that the disruption of NMJ guttering results from injury and regeneration, rather that dystrophin deficiency (9). Our finding that 2-week-old, pre-necrotic mdx muscles do not display fragmented NMJs is consistent with that interpretation. Thus, the decreased NMJ fragmentation in older NOS Tg/mdx muscles may reflect the function of NO in inhibiting mdx muscle damage by inflammatory cells (24). However, our further findings that muscle-derived NO contributes to AChR expression and clustering are not explicable by reductions in muscle injury and repair, and reflect a more direct regulatory function for NO in affecting synapse structure.

Several observations support a role for NO in modulating both AChR expression and clustering in vivo. The reductions of muscle NOS concentration in nNOS –/– mice, mdx mice (21,22), {alpha}-syn –/– null mice (11) and even in slow muscles of wild-type animals (25) were all associated with a decrease in AChR density at the post-synaptic membrane. In addition, the reduction of AChR densities in EDLs of mdx and {alpha}-syn –/– mice was partially reversed by expression of the NOS transgene. Furthermore, the significant increase in AChR density in NOS transgenic muscle was reversed by treatment with NOS inhibitor, which further supports a direct involvement of NO in promoting AChR clustering in vivo.

Although our data show a decrease in AChR clustering at the post-synaptic membranes of mdx mice, previous investigations have produced conflicting conclusions concerning whether mdx muscles have lower concentrations of AChR at the post-synaptic membrane. Electrophysiological findings have shown large reductions in miniature end-plate potentials (MEPP) in adult mdx mice, compared with controls (5,29), although one investigation reported only a statistically insignificant trend toward reduced MEPP in mdx muscles (2). Reductions in MEPP would be consistent with the reductions in AChR concentration at the post-synaptic membrane that are reported here. However, previous investigators have concluded that AChR concentrations at the post-synaptic membrane are not reduced in mdx muscle. These discrepant conclusions are attributable to the methods for measuring AChR concentration. In the investigations in which no reductions of AChR concentration at the mdx NMJ were reported, AChR concentration was determined by measuring radioactive decay in strips of muscles (29) or 1 mm slices of muscle (2) that had been incubated with 125I–BTX. However, those assays were unable to distinguish between synaptic and extrasynaptic AChRs, and defects in AChR clustering would be undetected. In the present investigation, fluorescent signal from FITC–BTX was measured only at the post-synaptic membrane, to reveal reduced AChR concentration and defective clustering at the NMJ in mdx and other NOS-deficient muscles.

Our findings show that increased NO production can promote AChR clustering that is stimulated through more than one signaling pathway. Although the signaling pathways through which laminin and VVA B4 induce AChR clustering show many similarities, they are not identical. For example, both ligands increase NOS activity and induce clustering through pathways that are dependent on NO (current investigation) and independent of MuSK or AChR-ß phosphorylation (30–32), but their effects on clustering are additive (33). The additive effect on clustering on myotubes stimulated with both ligands may reflect the ability of laminin to induce clustering by binding to either {alpha}7ß1 integrin (34) or {alpha}-dystroglycan (35,36), whereas VVA B4 binds {alpha}-dystroglycan (32,37,38), but is not known to bind {alpha}7ß1 integrin. Interestingly, clustering that is induced by agrin stimulation of myotubes is also reduced by administration of NOS inhibitors (39,40), but it occurs through a mechanism that involves MuSK and AChR-ß phosphorylation (41,42). Thus, the role of NO in promoting AChR clustering that is induced through at least three non-identical signaling pathways suggests that there may be overlap between NO-mediated signaling events induced by binding of each ligand. Potential events downstream of increased NO production that could lead to AChR clustering include a NO-stimulated increase in the expression of subsarcolemmal structural proteins that play a role in AChR clustering. For example, NO increases the expression of both talin and vinculin, which are part of the integrin complex that contributes to stabilizing the architecture of the post-synaptic membrane (43,44).

The large increase in mRNA levels for AChR-{alpha}, as well the significant increase in AChR-{alpha} concentration in muscle membrane pellets of NOS transgenic mice indicates that the increase in AChR-{alpha} concentration also reflects an increase in expression. Although the mechanism through which NO may increase AChR expression is not known, previous investigations suggest candidate pathways. For example, neuregulin or exogenous NO can elevate expression of c-fos and junB (45–47), and the neuregulin-induced expression of the epsilon subunit of the AChR requires c-jun expression (47). Similarly, c-jun N-terminal kinase (JNK) can be activated by either NO (48) or neuregulin (47), and JNK activation may be essential for neuregulin-induced expression of AChR (47). Those observations suggest the possibility that there may be overlap between signaling pathways that are mediated by NO and neuregulin that lead to elevated expression of AChRs. Previous investigators have also shown a decrease in AChR stability as a consequence of null mutation of DGC complex members (49), so that it is feasible that increases in NO may further elevate AChR concentration by increasing receptor stability, as well as by increasing expression.

The partial rescue of NMJ structure by expression of the NOS transgene in {alpha}-syn –/– EDL and quadriceps muscles (which express ß1-syntrophin), but not in {alpha}-syn –/– STM (which lacks ß1-syntrophin) indicates that expression of either {alpha}- or ß1-syntrophin is required for NO modulation of NMJ structure in vivo. Because NOS can bind either {alpha}- or ß1-syntrophin, the partial rescue of NMJ structure in {alpha}-syn –/– EDL and quadriceps could feasibly be attributed to NOS binding ß1-syntrophin at the post-synaptic DGC. This binding apparently requires high levels of NOS expression because NOS is not detectable by immunohistochemistry at the sarcolemma of {alpha}-syn –/– quadriceps muscles that do not express the NOS transgene (11). Partial rescue of the {alpha}-syn –/– EDL and quadriceps by NOS transgene expression could result from either NO-mediated events or processes affected by NOS–syntrophin interactions. However, two observations indicate that NO production rather than the physical binding of NOS to post-synaptic protein complexes provides the mechanism through which NOS affects synapse structure. First, our L-NAME treatment of NOS transgenic animals reversed the increase in AChR clustering and NMJ size that were caused by transgene expression. In addition, exogenous NO donors can induce increases in AChR expression and clustering that may contribute to the rescue of NMJ structure.

The findings presented here may also provide insights into the mechanisms through which increased muscle use can lead to adaptations in neuromuscular transmission and NMJ structure. Numerous reports have indicated that regular muscle use, such as during frequent exercise, causes adaptations in the structure and electrophysiology of NMJs (50–53). These adaptations may be relevant to the NO-mediated increases in AChR expression and density, and NMJ size, because muscle loading increases both NOS expression and activity (54). In the present investigation, we found that the area of the muscle fiber that was occupied by the NMJ was 25.5% greater in NOS transgenic mice than in non-transgenic controls. Intriguingly, regular exercise in rodents has been shown to increase NMJ area by 22 (50) and 23% (51). Similarly, regular exercise increased AChR concentration at the NMJ of rodents by 20% (52), compared with the 33% increase in NOS transgenic muscles, relative to controls.

Growing evidence shows that reductions of NOS expression can contribute broadly to the pathology of dystrophin-deficient muscle. Previous investigations have shown that NOS deficiency or mis-localization in the absence of {alpha}-syntrophin contributes importantly to defects in regulating peripheral blood flow following {alpha}-adrenergic stimulation (55,56), and that these vascular defects may promote the pathology of DMD (57). Other findings have shown that the loss of normal NO production by muscle causes increased inflammation of dystrophin-deficient muscle, and that the invading macrophages then promote muscle membrane lysis (24). The present findings which show that NOS deficiency can also lead to defects in synapse structure, further emphasize the important role of NO in muscle homeostasis, and illustrate the multiple cellular defects that can arise from its loss from muscle. Although these findings suggest that NO-based therapeutics could be valuable in addressing diverse defects that arise from loss of some proteins in the DGC, whether those therapeutics will be effective with systemic delivery of NO or would require muscle targeting of NO donors has not been explored.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Animals
All experimentations using animals were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and the University of California, Los Angeles Institutional Animal Care and Use Committee. C57BL/6J mice (wild-type), mdx mice [C57BL/10ScSn-Dmd(mdx)] and nNOS –/– mice (B6;129S4-NOS1tm1P1h/J) were obtained from The Jackson Laboratories (Bar Harbor, ME, USA). Transgenic mice with a muscle-specific, over-expression of nNOS (NOS Tg mice) were generated as described previously (24). Null mutant mice for {alpha}-syntrophin ({alpha}-syn –/– mice) were produced previously (11). Mdx mice were crossed with NOS Tg mice to produce dystrophin-deficient, NOS Tg mice (NOS Tg/mdx mice), according to previously described breeding strategy (24). NOS Tg mice that were null mutants for {alpha}-syntrophin (NOS Tg/{alpha}-syn –/– mice) were produced by the same breeding strategy. Null mutation for dystrophin was confirmed by ARMS PCR (58). Null mutation for nNOS or {alpha}-syntrophin was confirmed by western analysis of muscle extracts. Over-expression of nNOS was also confirmed by western blots of muscle extracts. Some NOS Tg mice were provided NOS inhibitor (L-NAME) in their drinking water at 0.5 mg/ml for 7 days prior to experimentation.

AChR distribution in vivo
The distribution of AChRs in wild-type and genetically-modified mice was assayed by labeling single muscle fibers with FITC–BTX. Mice were killed by exposure to carbon dioxide. Muscles were exposed by dissection and then immersed in 2% paraformaldehyde in phosphate buffered saline (PBS) after which single muscle fibers were dissected from the EDL, STM and SOL muscles. Individual, intact fibers were then incubated in FITC–BTX at 1 µg/ml in PBS for 1 hr, rinsed with PBS and mounted on glass coverslips. Images were obtained by epifluorescence microscopy using an Optronics CCD camera mounted on an Olympus BH2 microscope. Data were collected only from fibers in which the NMJs were viewed en face on the mounted fiber. At least five mice from each strain were analyzed, with a minimum of 10 NMJs per data set.

Measurement of the length of post-synaptic membrane crests
The crests of the folds of the post-synaptic membrane are highly enriched in AChRs and thereby provide the site of most AChR binding during synaptic transmission at the NMJ. We compared the length of the crests of the post-synaptic membranes of fibers from each strain of mice by collecting images of the BTX-labeled NMJs, and then using edge-finding software (Bioquant, R&M Biometrics, Nashville, TN, USA) to delineate the borders between AChR-rich crests and adjacent areas (Fig. 9). Stereological measurements were made of the lengths of post-synaptic membrane crests at each NMJ using an intercept point-counting technique with a Merz sampling grid (59).



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  		Figure 9. (A) AChR distribution and concentration at the post-synaptic membrane was measured in single fibers stained with FITC–BTX using en face views of the synapse. (B) AChR-rich post-synaptic membrane was delineated using edge-finding software. The fluorescence intensity within the delineated area was measured as an index of AChR density. The length of the AChR-rich crests was measured by stereological techniques to determine length of the post-synaptic membrane.

 
Measurement of post-synaptic membrane area
The dispersal of AChRs on the surface of each fiber was determined by measuring the surface area of each fiber that was occupied by each NMJ. Images collected from each BTX-labeled NMJ were analyzed by determining the simple, closed boundary with the smallest perimeter that would enclose all NMJ AChRs on the surface of each fiber, and then by measuring the enclosed area (Bioquant).

Measurement of AChR density at the post-synaptic membrane
The concentration of AChRs at the crests of the post-synaptic membrane was determined by measuring the average fluorescence intensity of BTX at the NMJ. Images that were used for analysis were all obtained under constant, manual exposure conditions. Exposure time was selected by determining empirically the length of exposure for the brightest field that was below saturation. Measurements were corrected for background signal by subtracting the signal obtained from a region of the slide where no muscle fiber was present. Images for which the boundaries between AChR-rich crests and adjacent areas were identified by edge-finding software were analyzed by measuring the area within the delineated boundaries, and the fluorescence intensity of FITC–BTX within that area (e.g. the fluorescence intensity density within the boundaries shown in Fig. 9). Relative AChR concentration was expressed as the total fluorescence intensity per total area delineated for each NMJ.

Measurement of relative AChR concentration in muscle membranes
Mice were killed by exposure to carbon dioxide, and then all hindlimb muscles were rapidly removed, and cleaned of fat and large pieces of connective tissue. The muscles were minced with a razor blade and homogenized (Sorvall Omnimixer) in 40 ml of buffer per gram of muscle (buffer: 50 mM Tris pH 7.5 containing 1 mM dithiothreitol, 1 mM ethylene diamine tetra-acetic acid, 1 mM benzamidine, 0.5 mM phenylmethylsulfonyl fluoride, and 1 µg/ml of aprotinin, 2 µg/ml leupeptin and 10 µg/ml soybean trypsin inhibitor). The homogenate was then filtered through several layers of gauze, and the filtrate was centrifuged at 12 000g for 15 min at 4°C. The supernatant was then centrifuged at 100 000g for 30 min at 4°C. The pellet obtained contained the particulate fraction that was enriched in vesiculated membranes, and the supernatant was enriched in cytosolic proteins. Protein in the supernatant fraction was concentrated by centrifugation through a 10 000 molecular mass cut-off cellulose membrane (Centriprep). The pellet fraction was solubilized by boiling in reducing sample buffer. Protein concentration in the pellet and supernatant fractions was measured using the technique of Minamide and Bamburg (60).

A 30 µg aliquot of each sample was electrophoresed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) (61) and the proteins in the SDS–PAGE gel were electrophoretically transferred to nitrocellulose before immunoblotting for nNOS (mouse anti-nNOS; Transduction Labs, Lexington, KY, USA) or for the {alpha}-subunit of the AChR (rat anti-AChR-{alpha}; Sigma, St Louis, MO, USA). Bands were visualized using enhanced chemiluminescence using a fluorescent chemiluminescence imaging system with which images were collected below saturation levels for the imaging system (Fluorochem 8900 Chemiluminescence Imaging System, Alpha Innotech, San Leandro, CA, USA).

Expression profiling
All hindlimb muscles were dissected from three adult mice of each of the following strains: NOS Tg, non-transgenic littermates of NOS Tg mice, NOS Tg/mdx mice and non-transgenic littermates (non-Tg/mdx mice) of NOS Tg/mdx mice. Muscles from the three mice of each strain were pooled, and RNA was isolated (62) and reverse transcribed using a SuperScript system (GIBCO BRL) with an oligo-dT primer and used to generate cRNA for expression profiling according to previously described procedures (24). Affymetrix microarrays (murine U74A gene chip) and software were used for analysis.

Immunohistochemistry
EDL, SOL, STM or quadriceps muscles were dissected from mice and then rapidly frozen in isopentane cooled in liquid nitrogen. Transverse, frozen sections of muscles were cut at 10 µm thickness, fixed in acetone and then labeled for nNOS (rabbit anti-nNOS; Serotec, Raleigh, NC, USA), syntrophin (SYN1351) (11), or AChR (FITC–BTX; Sigma). Appropriate, species-specific second antibodies were used.

Cell culture
C2C12 muscle cells in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS) were cultured on coverslips that had been coated with 2% gelatin in 50 mM sodium phosphate pH 7.2 containing 150 mM sodium chloride (PBS). When cells reached confluence, they were transferred to DMEM without FBS overnight to induce fusion. They were then returned to DMEM containing 10% FBS for 4–6 days prior to use in experiments.

Measurement of AChR clustering in vitro
C2C12 myotubes were incubated for 0, 1, 2, 6 or 24 h in 100 µg/ml of VVA B4 or with 200 nM laminin isolated from basement membrane of EHS mouse sarcoma (Sigma) in DMEM with 10% FBS (Sigma) to induce clustering of AChR. L-NAME was added at 200 µM to some preparations. Control preparations were incubated in DMEM with 10% FBS for the same period. At the end of incubation, the myotubes were rinsed two times with DMEM and then incubated with FITC–BTX diluted to 1 µg/ml in DMEM. Cells were then rinsed two times in PBS and then fixed for 10 min in 2% paraformaldehyde in PBS, before rinsing and mounting for microscopic observation.

The number of AChR clusters was measured by epifluorescence microscopy for a minimum of eight coverslips per treatment. For each coverslip, all clusters were counted in 112 fields that measured 3215 µm2 each, for a total of 0.36 mm2 sampled per coverslip. Data were then expressed as the number of clusters/mm2. In addition, the relative concentrations of AChR in clusters were compared by measuring fluorescent intensity of each cluster using a Bioquant digital imaging system.

Measurement of VVA B4 and laminin stimulated NO production in vitro
Myotube cultures were prepared and stimulated with VVA B4 or laminin for 1 h using the same conditions as used for stimulating AChR clustering. At the end of the VVA B4 or laminin stimulation period, DMEM was withdrawn from the cultures, and analyzed for NO concentration using a Sievers NO analyzer (Boulder, CO, USA). NO measurements were performed by reducing nitrites in the media to NO, and then reacting NO with ozone in a gas-phase chemiluminescent reaction.

Measurement of AChR surface expression in vitro
The relative concentrations of AChR on the surface of myotubes was compared by measuring the fluorescent signal intensity of FITC–BTX labeled myotubes. Fluorescence was measured in an 8 µm sampling circle on the surface of 100 myotubes that were >20 µm in diameter, but were otherwise sampled randomly. Images were collected as described for measurements of AChR density at NMJs of single muscle fibers, using the same procedures for background correction and controlling for saturation of signal. Myotube cultures that were used for measuring the relative concentrations of surface AChR were sampled at 0, 2, 6 or 17 h following incubation with 200 µM L-NAME, or with the following NO donors (Calbiochem, San Diego, CA, USA): sodium nitroprusside (SNP), SNAP, N-ethyl-2-(1-ethyl-2-hydroxy-2-nitrosohydrazino)-ethanamine (NOC-12) or NOC-18. NO donors were diluted in DMEM at 0.1, 0.5, 1.0 or 10.0 mM. Control cultures received DMEM only.


   ACKNOWLEDGEMENTS
 
Ms Katherine Wen provided excellent technical assistance. This work was supported by grants from the National Institutes of Health AR40343 (J.G.T.), AR47721 (J.G.T.) and NS33145 (S.C.F.) and the Muscular Dystrophy Association, USA.


   FOOTNOTES
 
* To whom correspondence should be addressed at: Department of Physiological Science, 5833 Life Science Building, University of California, Los Angeles, CA 90095, USA. Tel: +1 3102063395; Fax: +1 3108258489; Email: jtidball{at}physci.ucla.edu


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