Sarcoglycans are Enriched at the Neuromuscular Junction in a Nerve-Dependent Manner

doi:10.21203/rs.3.rs-4876243/v1

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Sarcoglycans are Enriched at the Neuromuscular Junction in a Nerve-Dependent Manner

https://doi.org/10.21203/rs.3.rs-4876243/v1

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Sarcoglycanopathies are heterogeneous proximo-distal diseases presenting severe muscle alterations. These diseases are caused by mutations in genes coding for one of the four sarcoglycan transmembrane proteins, which form the sarcoglycan complex (SGC). Little is known about the different roles of the SGC beyond the dystrophin glycoprotein complex (DGC) structural role. Here, we show that SGC proteins are enriched at the post-synaptic membrane of neuromuscular junctions (NMJs). Using a mouse model lacking the beta-sarcoglycan subunit, we describe for the first time that the loss of the SGC in the NMJ area results in alterations of pre- and postsynaptic membrane, as well as a significant reduction of membrane potential. Moreover, using different denervated wild-type mouse models, we demonstrate that enrichment of sarcoglycans within the NMJ occurs only after innervation, suggesting a nerve-dependent sarcoglycan expression. Altogether, our findings suggest that pathological decline should no longer be understood only in terms of sarcolemma damage but also in terms of sarcoglycans' participation in the NMJ. Henceforth, our work paves the way for the identification of new mechanisms involving sarcoglycans and new approaches for the treatment of sarcoglycanopathies.

Biological sciences/Neuroscience/Development of the nervous system/Synaptic development

Health sciences/Diseases/Neurological disorders/Neurodegeneration

Limb Girdle Muscular Dystrophy (LGMD) types R3, R4, R5, and R6, collectively known as sarcoglycanopathies, encompass a spectrum of genetic disorders characterized by significant heterogeneity that leads to progressive muscle weakness and reduced life expectancy (1–4). Currently, there is no cure for sarcoglycanopathies (5).

These conditions arise from mutations in genes encoding one of the four subunits of the sarcoglycan complex (α, β, γ, and δ), critical components of the dystrophin-associated glycoprotein complex (DGC) (6–8). Loss or dysfunction of any of the sarcoglycan subunits jeopardizes the whole sarcoglycan complex (9), rendering muscle fibers susceptible to damage and degeneration. However, there are conflicting data demonstrating that some residual subunits may exist in some patients with mutations in gamma or alpha sarcoglycan genes, indicating that incomplete complexes could be found in the fiber membranes (10, 11).

The primary role of the sarcoglycan complex is maintaining sarcolemma integrity within the DGC. However, the enormous heterogeneity observed among patients and even within muscles of the same patient suggests its involvement in other cellular processes (12, 13). In accordance with this argument, various proteins of the DGC, such as dystroglycan, dystrobrevin, syntrophin, and even dystrophin, have been found to have non-mechanical roles, including activation of nitric oxide pathways (14), and maturation and maintenance of the neuromuscular junction (NMJ) (15–18).

A prominent feature of the neuromuscular junction is the dense clustering of acetylcholine receptors (AChRs) at the postsynaptic membrane (19). In embryonic stages, AChRs are initially distributed evenly across the membrane of the newly formed fibers. With innervation, the pentameric receptor aggregates at sites of nerve contact and stabilize; replacing the gamma subunit by the epsilon subunit (20–22). This nerve-induced process involves multiple molecules and signaling pathways such as Agrin-Lrp4-Musk pathway which includes Agrin, Lrp4, Musk, Dock7, and rapsyn (23, 24). The maturation-clusterization process can also be observed in adult muscle due to the daily turnover of the fibers or after acute damage (25). Following acute damage, muscle fibers undergo regeneration and this process involves a meticulous net of signals, with the final aim to restore the muscle homeostasis, and to reinnervate the newly formed muscle fibers (26). When the process of fiber degeneration /regeneration becomes chronic, muscle fibers loss their ability to contrast the damage, starting a process of degeneration and atrophy (25, 27, 28). Interestingly, accumulating evidence indicates that the nerve-muscle crosstalk may be bidirectional with muscle damage contributing to nerve transmission decline, and nerve damage contributing to muscle atrophy (29).

However, our knowledge of NMJ alterations during chronic muscle diseases comes mainly from the mdx model, where Dystrophin has been observed at the muscle endplate, in particular in the depths of the junctional folds, where it binds voltage-gated Na + channels and determines the length of synaptic folds (30). In this mouse model, severe alterations such as arborization of nerve terminal, fragmentation of postsynaptic membrane and transmission failure exacerbate muscular decline (31–33). Regarding sarcoglycans, only a few early studies have suggested that these proteins may be located at the NMJ (34, 35), but there is paucity of data regarding the impact of sarcoglycans deficiency on neuromuscular communication. Recently Fornetti et al. (36) and Zhao et al. (37), have established a link between alpha-sarcoglycan and the NMJ, pointing to the importance of this complex for nerve-muscle dialogue.

In this work we investigated the presence of sarcoglycans around the AChR-covered area. Moreover, we investigated whether SGC disruption alters the NMJ structure and whether nerve transmission interruption could disrupt the SGC signal.

This study aimed to expand our knowledge of the sarcoglycan complex and its functions in muscle. We lay the foundation for future studies of sarcoglycans’ involvement in other molecular pathways, which could help us better explain the pathophysiology and phenotypic diversity among patients or among muscles of the same person affected by sarcoglycanopathy.

1. Sarcoglycan subunits are present at the NMJ.

To understand whether sarcoglycan subunits participate in the NMJ, we decided to search for their presence in available online databases. We re-analyzed the Millay’s lab MyoAtlas dataset (https://research.cchmc.org/myoatlas/), which provides single-nucleus skeletal muscle gene expression data (38). We observed RNA expression of the four sarcoglycan subunits (alpha, beta, gamma, and delta), in nuclei belonging to the neuromuscular junction cluster (Fig. 1A-C). Next, it was analyzed the signal intensity of sarcoglycan proteins along the sarcolemma (Fig. 1D). We identify an enrichment of the four sarcoglycan subunits signal, colocalizing to alpha-bungarotoxin.

2. The lack of beta-sarcoglycan alters NMJ morphology.

We performed whole mount immunofluorescence on isolated WT fibers to better understand the location of sarcoglycan subunits at the NMJ. As shown in Figure 2A, protein localization in the NMJ was observed for the four analyzed subunits. Next, using the beta-sarcoglycan KO mouse model (39) we observed a reduced expression or absence of the other sarcoglycan subunits at the NMJ. Notwithstanding, the gamma-sarcoglycan subunit was still detectable at the NMJ despite the lack of beta-sarcoglycan, suggesting its independent clustering dynamics compared to the others (Fig. 2A and Suppl. Fig. 1A). Of note, the AChR remained detectable despite sarcoglycans absence. On the other hand, in WT muscle, the alpha sarcoglycan signal was colocalized with AChR signal while the other subunits are close to AChR signal without overlapping it (Suppl. Fig. 1B).

Furthermore, we observed a reduction of dystrophin at the NMJ level (Fig. 2B), suggesting that the loss of beta-sarcoglycan may compromise other members of DGC. To further characterize this postsynaptic destabilization in the beta-sarcoglycan KO mouse model, we studied the morphological parameters of AChRs distribution in the endplate. We observed statistically significant increases of endplate area (Fig. 2C) and AChR fragmentation compared to WT (Fig. 2D), but no differences in compactness (Fig. 2E). Contrary to what was described in previous studies (40) we observed a similar amount of subsynaptic nuclei between WT and KO (Fig. 2F).

Additionally, despite no denervation being observed in Sgcb KO muscle sections, as shown by the high rate of colocalization between BTX and neurofilament (Suppl. Fig. 1C), axon terminals were deeply affected, showing an aberrant arborized morphology of nerve terminals (Fig. 2G). Altogether, these results demonstrate that the lack of beta-sarcoglycan protein results in morphological destabilization of both pre- and postsynaptic structures.

3. The lack of beta-sarcoglycan impacts membrane potential of muscle fibers.

To understand whether morphological changes in the NMJ could be associated to changes in AChR functionality, we recorded ACh-evoked channel activity on WT and Sgcb KO muscles. We used isolated fibers of flexor digitorum brevis (FDB), a glycolytic fast twitch distal muscle from 4-8 weeks old mice (Fig. 3A). Synaptic regions were eye identified, and openings of synaptic AChR-channels were characterized by the channel conductance, open duration and closed time of unitary events. In both strains, openings of only one channel population were detected (Fig. 3B and 3C). All functional parameters were similar in WT and KO fibers (Fig. 3D) and compatible with the openings of AChR-channels containing the epsilon subunit (22). In several fibers, recordings were also performed in non-synaptic regions, observing only occasional openings of AChR-channels (data not shown). These functional observations indicate that fibers were non denervated. However, the plot of unitary current of each single fiber vs. pipette potential showed a statistically significant reduction of the current reversal potential in Sgcb KO fibers (Fig. 3E).

Additionally, we found that the level of intracellular Ca2+ in another distal muscle such as TA, was slightly but significantly increased in Sgcb KO mouse fibers (Fig. 3F) despite not-significant differences on damaged fibers were observed histologically (Suppl. Fig. 2A). On the other hand, upregulation of AChR transcription levels was detected (Fig. 3G). We observed a significant increase of the alpha and gamma AChR subunits compared to WT. By contrast, the transcript level of the epsilon subunit, replacing the gamma subunit in adult innervated fibers, was similar to WT (Fig. 3G). Of note, based on patch-clamp experiments, no opening of AChR-channels containing the gamma subunit were detected in synaptic or extra-synaptic regions. In any case, the upregulation of AChRs transcripts was concomitant with a transcriptional upregulation of Agrin-Lrp4-Musk pathway (Fig. 3H), but not of choline acetyltransferase levels (Suppl. Fig. 2B). The upregulation pattern in Sgcb KO conditions was not observed during regeneration process after acute damage (Suppl. Fig. 2C and 2D).

4. Nerve damage disrupts the SGC at the NMJ.

To further understand the relation of sarcoglycans with NMJ structure and function, we investigated whether sarcoglycan enrichment within the NMJ structure was altered upon perturbations or damage to the nerve. We used two different mouse models of nerve damage: sciatic nerve crush and sciatic nerve cut (Fig. 4A) (41). These models allowed us to simulate reversible or chronic damage, respectively. We analyzed sarcoglycan expression and distribution by whole-mount immunofluorescences at time 0 (before the damage), at 7 and 30 days after nerve crush, or 30 days after nerve cut (Fig. 4A). In both cases, we observed a severe reduction of sarcoglycan subunits when the muscle was denervated: 7 days after crush or 30 days after cut (Fig. 4B). The signal was reestablished to basal levels 30 days after crush when the muscle was reinnervated. No alterations were observed in the contralateral legs (Suppl. Fig. 3A).

Along with the loss of sarcoglycan subunits, we observed reduced endplate size and increased AChR signal fragmentation but not changes in compactness (Fig. 4C). Interestingly, the expression of the gamma sarcoglycan subunit, which retained some level of expression in the absence of beta-sarcoglycan (Fig. 2A and Suppl. Fig. 1A), was dependent on nerve signals, as were the other subunits. We observed no changes on morphological parameters at the contralateral leg (Suppl. Fig. 3B). Altogether, these results suggest that the enrichment of the sarcoglycan complex at the NMJ is dependent on nerve signals.

5. Sarcoglycans enrichment at the NMJ is secondary to AChRs-spot formation.

To further investigate whether sarcoglycan enrichment at NMJs is due to the nerve presence, we used an in vitro system. C2C12 cells were cultured and differentiated into myotubes (Fig. 5A). We then induced AChR clusters in myotubes, treating them with agrin (Fig. 5B) (42). As expected, an increase in AChR clusters was observed after treatment (Fig. 5C). However, no accumulation of sarcoglycan subunits at the sarcolemma and in AChR-spots show was detected. While alpha, beta and delta sarcoglycan subunits maintain similar levels of expression, the gamma subunit was widely downregulated (Fig. 5D). Altogether these results indicate that no enrichment of sarcoglycan subunits at the area of AChR clustering occurs in the absence of nerve.

Sarcoglycans are transmembrane proteins mainly expressed in muscle fibers. Even though their main role has been primarily related to support of the sarcolemma within the DGC, the expression of some subunits has also been linked to the NMJ (36, 37). However, no studies have characterized their presence as a complex in the postsynaptic structure until now.

In this study, we show for the first time that the entire SGC is enriched at the NMJ. We observed that the absence of the complex alters the morphology of pre and postsynaptic membranes in the NMJ and provokes a reduction of the membrane potential even in distal less affected muscles. Finally, using different models of denervation we provide evidence that sarcoglycans enrichment specifically requires nerve-derived signals other than agrin, suggesting that their presence might be linked to the presence of the nerve, and possibly to some unknown specific role(s).

First, on WT mouse we observed an enrichment of sarcoglycan signal corresponding to the AChR clusterization sites. Furthermore, using a Sgcb KO mouse model, we observed differences between the SGC in the sarcolemma and the SGC at the NMJ. Whereas in the sarcolemma, the loss of beta sarcoglycan subunit results in the loss of all subunits, at the NMJ, the gamma subunit expression was preserved (Fig. 2A). This finding suggests that the gamma subunit may follow an independent clustering pathway from the rest of the complex. Additionally, observations of no defects in NMJ in the gamma-sarcoglycan KO mouse model (43), further support the idea that the gamma subunit may move to NMJ using alternative pathways.

Next, in the absence of SGC induced by beta-sarcoglycan deficiency, we observed morphological changes in pre and postsynaptic elements of the NMJ, including arborization of nerve terminal and enlargement and fragmentation of postsynaptic membrane. These findings, reported for the first time in sarcoglycanopathies, have been previously described in mdx mice (40, 44) and in chimeric mice lacking dystroglycans (45). However, it remains unclear whether they are a secondary effect of sarcolemma damage or a direct effect of the degeneration/regeneration process (46). In spite of limited structural damage, we report depolarization of resting membrane potential and increased levels of intracellular Ca2+, possibly secondary to sarcolemma damage. On the other hand, we observed increased transcriptional levels of some AChR subunits together with increased levels of the Agrin-Lrp4-MuSK pathway, but no changes in acetylcholine transferase (ChAT) protein presence. These findings, observed in mdx as well (33), may suggests an activation of the regeneration process in Sgcb KO muscles, although we observed no upregulation of AChR subunits or Agrin-Lrp4-MuSK pathway during regeneration process generated after Ctx injection (Suppl. 2C and 2D). Therefore, we believe that one possible explanation of the AChR upregulation could lie in some compensatory mechanism related to the subsynaptic nuclei.

To further investigate the signals that trigger SGC accumulation in the NMJ, we conducted experiments in WT mice with partial or total denervation of the sciatic nerve and in vitro experiments. From these experiments, we observed that after nerve crush or cut, sarcoglycans disappeared from the postsynaptic part of the NMJ, but persisted in the sarcolemma. These findings, together with our in vitro experiments performed in the absence of nerve where no accumulation of sarcoglycans in the area of AChR clusters occurred, identify the nerve as an essential upstream element for sarcoglycan complex enrichment at the NMJ.

Despite several limitations, such as not being able to identify the type of subsynaptic cells expressing enhanced levels of sarcoglycans or whether the arborization of the nerve terminal is related to declined neurotransmission, our study suggests an unknown nerve-dependent link between the NMJ and the SGC. This link could be among the reasons why muscle pathologies commonly increase their degeneration rates after patients lose ambulation. However, further research is needed to unravel the possible role of the sarcoglycan complex at the NMJ and its involvement in new molecular pathways. The multiple glycosylation and phosphorylation sites of all sarcoglycan subunits(47) might help validate or refute some hypotheses.

In conclusion, our work opens new avenues for exploring the role of SGC in the NMJ and its contribution to sarcoglycanopathies focusing on the nerve’s influence on muscle disease. Understanding the specific role of the sarcoglycan complex at the NMJ could lead to innovative therapeutic approaches targeting this critical synaptic interface.

Bioinformatic analysis

The post-natal (P) 21-day WT dataset from Petrany and colleagues (38) was obtained by subsetting the fully processed scMuscle compendium Seurat object from McKellar et al., available for download on Dryad (48). Data were normalized using Seurat’s SCTransform function (Seurat v5) (49). Dimensionality reduction was performed through Principal Component Analysis (PCA) on top variable features, with npcs = 30. FindNeighbors, FindClusters, and UMAP embedding functions were based on the top 30 PCs (Principal Components) of the PCA. We identified 12 clusters using FindClusters with parameters resolution = 0.9. After clustering, cell populations were annotated using the “FindAllMarkers” function according to well-known lineage markers, resulting in 9 meta-clusters. The annotated dataset was used to generate the dotplot in Fig. 1B and the FeaturePlot in Fig. 1C.

Denervation

TA muscles from 4-week-old female WT C57BL/6J mice were used for this experiment. Briefly, mice were deeply anesthetized with a mixture of Rompun (Bayer, 20 mg/mL; 0.5 mL/kg) and Zoletil (100 mg/mL; 0.5 mL/kg). For crush injuries, the sciatic nerve was compressed at three different points for 5 seconds. For cut injuries, approximately 3 mm of the sciatic nerve was removed, followed by wound closure with tissue adhesive (Vetbond). Buprenorphine (0.05–0.1 mg/kg) was administered subcutaneously prior to the animals regaining consciousness (41).

Immunofluorescence

TA muscles from WT C57BL/6J and Sgcb KO mice (B6.129-Sgcbtm1Kcam/2J) females, aged 4 to 8 weeks, were used to obtain cryosections or whole mount preparations of entire fibers. Cryosections were permeabilized with 100% acetone, while fibers were fixed in 4% PFA (MilliporeSigma, P6148) at room temperature. A blocking solution containing 4% BSA (MilliporeSigma, A7030-100G) was applied to both types of preparations. Primary antibodies were applied overnight at 4°C: anti-Sgca (Ab189254), anti-Sgcb (Ab315386), anti-Sgcg (Nova Castra 6091787), and anti-Sgcd (kindly provided by Dr. Sandonà). Secondary antibodies goat anti-rabbit Alexa Fluor 594 (A11011) and goat anti-mouse Alexa Fluor 594 (A21235) were then applied. Alpha-Bungarotoxin coupled to Alexa Fluor 488 (B13422) was used to identify AChR presence. Counterstaining was performed with DAPI (Thermo Fisher Scientific, D1306). All images were acquired using confocal microscopy with z-stack intervals of 1 µm. Images were subsequently analyzed using ImageJ software.

Morphological analysis

Morphological analysis consisted in the assessment of endplate area, fragmentation, compactness and subsynaptic nuclei clusterization. Additionally, colocalization of AChR signal with sarcoglycans was also analysed. All analyses were performed following standard procedures described elsewhere (50).

Muscle fiber preparation and cell-attached patch-clamp recordings

Flexor digitorum brevis (FDB) muscles from WT and Sgcb KO mice were dissected from the hindlimbs of female mice aged 4–8 weeks, which were sacrificed by cervical dislocation. The FDB muscles were incubated with Type I collagenase (3 mg/mL, Sigma) for 30 minutes at 37°C in Minimum Essential Medium (MEM) (Gibco, Thermo Fisher Scientific, #11095-080). After equilibrating in Ca2 + free suspension culture MEM (Gibco, #11380-037) for 30 minutes at room temperature, the muscles were gently triturated until to obtain single fibers. Recordings were performed on fibers imaged by a phase-contrast microscope (Axioskop 2 FS, Zeiss, Jena, Germany). Patch pipettes were filled with a solution containing (in mM, all from Sigma Aldrich, St. Louis, MO, USA): NaCl 140, KCl 2.8, CaCl2 2, MgCl2 2, glucose 10, HEPES/NaOH 10, pH 7.3, plus ACh (100 nM). The resistance was 4–6 MΩ. Single-channel currents were recorded at room temperature (23–26°C) using a low-noise Axo-patch 200B amplifier (Molecular Devices, San Jose, CA, USA) in cell-attached mode. Data were sampled at 25 kHz, digitally filtered at 5 kHz (Gaussian filter), and analyzed using pClamp 9.2 (Molecular Devices). Only single openings (100–3000 at each pipette potential in every patch) were used to determine the channel slope conductance and duration. Measurements were performed at the same pipette potential at the beginning and end of each recording to ensure membrane potential stability. Slope conductance was calculated only if the membrane potential was stable and unitary events were recorded at least at three different pipette potentials. The kinetic properties of ACh-evoked events were compared at an estimated membrane potential of − 90 ± 10 mV, assuming a reversal potential of 0 mV. Histograms of open times were fitted with sums of exponential components using a non-linear algorithm (pClamp 9.2), as required.

RNA extraction and real time PCR

Total RNA was extracted from frozen female tibialis anterior muscles using TRIzol Reagent (Life Technologies, 15596018) following the manufacturer’s protocol. The RNA was then reverse transcribed using the High-Capacity cDNA Reverse Transcription Kit (Life Technologies, 4368814). For quantitative PCR (qPCR), premade TaqMan assays labeled with 6-Carboxyfluorescein (FAM) (Thermo Fisher Scientific) were used.

Intracellular calcium levels determination

INDO-1AM (Invitrogen, I1226) was reconstituted in high-quality, freshly opened DMSO to a concentration of 1 mM. For experiments, it was used at a final concentration of 100 µM in Ca2+/Mg2+-free PBS (phosphate-buffered saline). After reconstitution, the solution was protected from light and stored at − 20°C to prevent freeze-thaw cycles. Samples were washed three times with Ca2+/Mg2+ -free PBS before slowly adding INDO-1AM along the muscle fibers. The samples were then incubated for 30 minutes at 37°C. Following incubation, the samples were washed three times with Ca2+/Mg2+ -free PBS and analyzed using confocal microscopy. Detection was based on the dual emission properties of INDO-1, which shifts from 475 nm in Ca2+ -free media to 400 nm when saturated with Ca2+(51). All analyses were conducted using Zeiss Confocal Software (Zen 3.0 Blue Edition).

Western Blot analysis

Protein extraction and western blotting were carried out using standard techniques as detailed in Supplementary Figure S2.

In vitro cultures

C2C12 were cultured until differentiation following standard proceduresand then, stimulated with agrin (RD Systems, 550-AG) following the procedure described in Proietti et al. (52)

Statistical analysis

All statistical analyses were conducted using Prism version 9.0 (GraphPad Software, San Diego, CA, USA). Data are presented as the mean ± SD. Statistical significance was assessed using paired Student’s t-test, one-way ANOVA, as specified in each figure legend. A p-value of < 0.05 was considered statistically significant (∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001), while p-values ≥ 0.05 were considered not significant.

Study approval

All experiments in this study were conducted in accordance with protocols approved by the Italian Ministry of Health, Rome, Italy, and the ethical regulations of the University of Rome 1 “Sapienza” (Rome, Italy).

Conflict of interests:

The authors have declared that no conflict of interest exists.

Author contributions:

M.G., B.C., M.C., V. R., M.S.P. and F.G performed all experiments. C.D. performed the bioinformatic analysis. B.L., F.G, and M.B supervise and edit the work. M.B., L.M. and C.S.R. design the experiments.

Acknowledgment:

We want to thank the GFB Foundation for financially supporting this work. We would also like to acknowledge Dr. Egle Destefano for her support and valuable discussions, as well as Dr. Doriana Sandonà for kindly providing us with a self-manufactured delta-sarcoglycan antibody.

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Supplementary1.tif
Supplementary 1 A) Representative images of immunofluorescent transversal sections of Sgcb KO female’s Tibialis Anterior, stained with sarcoglycans (red), AchRs (green) and Dapi (white). Scale bar 100 μm. Associated histogram measurements of AchR and sarcoglycans colocalization signal at NMJ. n=3. B) Scheme of co-localization measurement; Sarcoglycan signal (white) and AchR perimeter (yellow) and histogram of co-localization measurements on WT fibers at 1 month. n=9. C) Immunofluorescent transversal sections of WT and Sgcb KO female’s Tibialis Anterior muscles, stained with Synaptophysin (red), BTX (green), Caveolin (cian) and Dapi (white). Scale bar 100 μm. Associated histogram of Syp and BTX double positive signal, associated to the immunofluorescence. n=3. One-way ANOVA for figures A and B, T.test for figure C. Significance was considered as *p=0.05; **p=0.01; ***p<0.001.
Supplementary2.tif
Supplementary 2 A) Representative images of immunofluorescent transversal sections of WT and Sgcb KO female’s Tibialis Anterior stained with IgG (red), Laminin (green), and Dapi (White). Scale bar 1000 μm. Associated histogram of IgG positive fibers expressed in fold increase. n=3. B) Western blot analysis was performed as described by Honda and colleagues (9). Primary antibodies used were anti-Choline Acetyltransferase (Synaptic Systems, 297015), and Gapdh (Cell Signaling, 2118L). Histogram related to the western. T.test. No significative differences. n=5. C) RNA levels of AChRs subunits performed in total muscle TA at 3, 7 and 15 days after Ctx damage and contralateral leg. D) RNA levels of Agrin pathway of the same muscles at 3, 7 and 15 days after Ctx damage and contralateral leg. Hprt was used as internal control. n=3. All analyses were made using T.test, *p=0.05; **p=0.01; ***p<0.001.
Supplementary3.tif
Supplementary 3 A) On the left, immunofluorescence whole mount preparations of contralateral Tibialis Anterior fibers of damaged (Crush and Cut) WT mouse at, 7 days after crush injury (reversable denervated), 30 days after crush injury (reinnervated) and 30 days after cut injury (denervated). The staining was made with antibodies against each Sarcoglycan subunit (red), BTX (green) and Dapi (blue). Scale bar 20 μm. On the right, histogram of sarcoglycan subunits intensity at endplate area. n=4. B) Morphometric parameters of endplate terminal; endplate area (left), fragmentation (centre) and compactness (right). n=11. One-way ANOVA, *p=0.05; **p=0.01; ***p<0.001.
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Sarcoglycans are Enriched at the Neuromuscular Junction in a Nerve-Dependent Manner

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