Advanced Diagnostics and Biomarkers – PLOS Currents Muscular Dystrophy Wed, 17 Oct 2018 20:45:27 +0000 en-US hourly 1 Benefits of Prenatal Taurine Supplementation in Preventing the Onset of Acute Damage in the Mdx Mouse Fri, 22 Sep 2017 11:00:30 +0000 Introduction: Duchenne Muscular Dystrophy (DMD) is a debilitating muscle wasting disorder with no cure. Safer supplements and therapies are needed to improve the severity of symptoms, as severe side effects are associated with the only effective treatment, corticosteroids. The amino acid taurine has shown promise in ameliorating dystrophic symptoms in mdx mice, an animal model of DMD, however little work is in 21-28 (d)ay animals, the period of natural peak damage. 

Methods: This study compares the effect of prenatal taurine supplementation on tibialis anterior (TA) in situ contractile function, histopathological characteristics and the abundance of Ca2+-handling as well as pathologically relevant proteins in non-exercised mdx mice at 28 and 70 d.

Results: Supplementation elevated TA taurine content by 25% (p<0.05), ameliorated in situ specific force by 60% (p<0.05) and improved histological characteristics in 28 d mdx mice; however no benefit was seen in 70 d mice, where background pathology was initially stable. Age specific effects in SERCA1, calsequestrin 1 (CSQ1), CSQ2, utrophin and myogenin protein abundances were seen between both 28 and 70 d mdx and mdx taurine-supplemented mice.

Discussion: Considering these findings and that taurine is a relatively cost effective, readily accessible and side effect free dietary supplement, we propose further investigation into taurine supplementation during pregnancy in a protective capacity, reminiscent of folate in the prevention of spinal bifida.



Duchenne muscular dystrophy (DMD) is a fatal, X-linked neuromuscular disorder affecting approximately 1:3500-6000 live male births1,2. DMD is caused by a mutation in the dystrophin gene which results in the absence of the cytoskeletal protein dystrophin. The dystrophic pathology is most notably characterised by progressively debilitating muscle weakness, as cumulative bouts of muscle degeneration exhaust regenerative capabilities and healthy myofibers are replaced by non-contractile, fat and fibrotic tissue. Ultimately this loss of dystrophin results in the loss of ambulation, respiratory complications, and lethal cardiac events by approximately 30 years of age3,4. There is currently no cure for DMD, and while genome editing using CRISPR/Cas9 technologies are showing great promise for future treatment5 there remains a persistent need for safer alternatives to current treatments utilising steroids1, which present the patient with a host of adverse side effects.

Dystrophin is part of a network of proteins known as the dystrophin-glycoprotein complex (DGC) that anchor the contractile machinery of a myofiber to the extracellular matrix. Whilst fundamentally this provides the muscle with stability during contraction there is increasing evidence that dystrophin, as a part of the DGC, also plays a role in cell signalling. Therefore while the precise mechanisms by which dystrophin deficiency leads to the muscle weakness seen in DMD remains unclear; it is clearly multi-faceted in nature6. The most common animal model of DMD is the dystrophin deficient mdx mouse, which has a premature stop codon in exon 23 of the dystrophin gene7. In this model, as well as other animal models, altered Ca2+ homeostasis; increased reactive oxygen species; inflammation and membrane destabilisation have all been implicated in the pathology of the disease6.

Importantly, the mdx mouse does not experience the progressive muscle wasting that leads to death in DMD patients8,9. Rather mdx mice experience an acute onset of hindlimb muscle necrosis from 21-28 (d)ays, at which time they most closely represent the severity of muscle pathology seen in DMD, before stabilising into low grade but persistent muscle damage into adulthood (>10 weeks). The up-regulation of utrophin, the structural homolog of dystrophin, as well as the cessation of growth and exercise avoidance behaviours all contribute to the milder phenotype observed in adult mdx mice9. Critically, however, DMD patients exhibit a more severe dystropathology, and whilst in response to initial bouts of damage and/or necrosis DMD patients experience myogenesis and regeneration, in contrast to the mdx mouse, this persists for years as opposed to weeks. The relatively stable pathology found in adult mdx mice can be exacerbated with damaging exercise protocols, and although this can be a source of inconsistency between studies provides a useful platform for the investigation of therapeutic interventions. However, the adult model does not allow the investigation of interventions that would act in a preventative or protective capacity to the acute phase of muscle degeneration, such as that observed naturally in the 21-28 d mdx mouse. Preventative strategies aimed at reducing the onset of degeneration, as opposed to those attempting repair when damage is overwhelming may show promise in alleviating dystrophic symptoms if the disease is diagnosed early.

Taurine, a sulphur containing amino acid, is found ubiquitously throughout the excitable tissues of the body where it is associated with healthy muscle function, differentiation and growth (For review see10,11). It is available commercially as a dietary supplement and is widely used as an ergogenic aid12. Of note, its use has no documented obvious side effects when supplemented in humans12. In healthy animals and mdx mice, taurine supplementation has been found to improve Ca2+ handling, membrane stabilisation, oxidative stress and inflammation both at rest and following damaging exercise protocols13,14,15. Recent investigations reported similar muscle taurine content in juvenile mdx mice at most ages (e.g. 18, 22, 36 and 42 days) compared to WT mice, although reduced in 28 d mdx mice13,16,17. These findings support that taurine levels vary with age in the growing mdx mouse. It appears that during the onset of mdx pathology taurine supplementation is efficacious at preventing the myofiber necrosis and inflammation in juvenile mdx mice if elevated early13. Of relevance, normal (non-dystrophic) mice deficient in the taurine transporter (TauT), as well as those with dramatically reduced intramuscular taurine content, experience dystrophic symptoms including compromised muscle strength, susceptibility to fatigue, and developmental abnormalities18,19. Taurine supplementation has been previously studied in vivo and in vitro in mdx mice, typically following chronic exercise protocols, finding both supplementation and direct application to muscle preparations effective at improving muscle health and force development16,20.

This study aims to investigate the efficacy of taurine at ameliorating dystrophic symptoms in the mdx mouse at two distinct pathological stages (28 and 70 d). Taurine was supplemented prenatally and throughout the life of animals to ensure the maximum effect of the amino acid. This study addresses the importance of age as a key consideration for pathological relevance when screening for therapeutic supplements, as well as when investigating biochemical and physiological properties of muscle from mdx mice. Our results show that at 28 d, during the onset of acute muscle damage, taurine was effective at increasing muscle strength and improving visual muscle health but shows no benefit to 70 d mdx mice where the background pathology is initially stable.

Materials & Methods

2.1 Animals and supplementation

All procedures in this study were approved by the La Trobe University Animal Ethics Committee (AEC 12-31, 13-48). Only male mice were used for experiments. A total of 35 mdx and 24 wild-type (WT, C57/BL10ScSn) were used. Experimental animals were bred at the La Trobe Animal Research and Teaching Facility using breeding pairs obtained from the Animal Resource Centre (Western Australia, Australia). The offspring of WT and mdx mice had access to standard rodent chow, water ad libitum and were utilised for experimentation at either 28 ±1 or 70 ±1 days of age. Mdx taurine (mdx tau) breeders and subsequent offspring were supplemented with continuous access to taurine (2.5% wt/vol) enriched drinking water, with breeders beginning supplementation at least two weeks prior to mating. This dosage of supplementation has been demonstrated previously to elevate skeletal muscle taurine content by up to 40% when given directly to rats21. The number of animals used in the various experiments sometimes differ due to sedation issues that occurred during in situ experimentation. This required the animal to be immediately culled.

2.2 Muscle Dissection

Mice were anesthetized with an initial intraperitoneal injection of 10ul.g-1 Nembutal (Sodium Pentobarbitone) and kept unresponsive for the duration of the experiment with supplementary doses (10% initial volume). The mouse was weighed, placed on a 37oC heated pad and the right leg was skinned to the waist taking care not to damage the fascia. Throughout the experiment, warmed physiological saline (0.9%) was applied to exposed muscle tissue. The distal tendon of the tibialis anterior (TA) muscle was isolated and secured with both a top and bottom knot as close to the myotendinous junction as possible using 5.0 surgical thread (Ethicon, Johnson & Johnson, NSW, Australia). The distal tendon was then severed and the TA dissected free from the surrounding tissue keeping nerve and blood flow intact at the tibia below the tibial plateau. The sciatic nerve was exposed by gently parting the midline of the biceps femoris muscle above the knee joint. The mouse was secured on the heated platform (37oC) of an in situ contractile apparatus (809B in situ Mouse Apparatus, Aurora Scientific, Ontario, Canada) with a pin behind the patella tendon and a foot clamp. The distal end of the TA was tied firmly to a lever arm attached to an isometric force transducer. The sciatic nerve was stimulated by two field stimulating platinum electrodes coupled to an amplifier.

2.3 Contractile Protocol

The TA was contracted via square wave (0.2 ms) pulses at 10V from the stimulator (701C stimulator, Aurora Scientific, Ontario, Canada). Forces were converted to a digital signal and recorded by Dynamic muscle analysis 611A (DMA) (Aurora Scientific, Ontario, Canada). Optimum muscle length (Lo) was first determined by eliciting twitch contractions at incrementally adjusting muscle length with a micromanipulator until a repeatable maximum peak twitch force (Pt) was obtained. Muscle length at Lo was measured with precision digital callipers from the beginning of the distal tendon to the insertion of the TA at the base of the knee. Subsequently the TA was stimulated at 100 Hz tetanic contraction, followed by a 2 min rest interval and then twitch contraction. Comparable twitch forces pre and post 100 Hz stimulation indicated that the knots were both secure and unlikely to slip during the remaining protocol. If a decrease in twitch force was observed the muscle was incrementally tensioned and stimulated between 2 min rest intervals until Pt was re-established. To establish the force frequency relationship the TA was stimulated for 500 ms at 10, 20, 30, 40, 50, 80, 100, 150 and 200 Hz with a 2 min rest interval in-between. Maximum isometric tetanic (Po) force was determined from the largest force produced during the force-frequency stimulation protocol. Maximum tetanic specific force (sPo) was determined as force per cross sectional area (CSA) as described by Lynch (DMD_M.2.2.005). Briefly, optimum fiber length (Lf) was first determined by multiplying Lo by the predetermined TA length to fiber length ratio of 0.622 and utilising the formula sPo = Po x (muscle mass/Lf x 1.06).

Following the force frequency protocol, the TA was allowed a 2 min recovery prior to commencement of the fatigue protocol. Fatigue was induced by stimulating the TA for 500 ms at 60 Hz every second for 180 s. At the completion of the fatigue protocol, recovery was assessed by stimulating the muscle for 500 ms at 60 Hz following 1, 2 and 3 min recovery periods.

Immediately following the contraction protocol, the mouse was removed from the apparatus and the contractile TA dissected, blotted clean on filter paper (Whatman No.1) and weighed, before being embedded in O.C.T. compound (Tissue-tek) and snap frozen in liquid nitrogen cooled isopentane (2-methylbutane, Sigma Aldrich). The contralateral TA was also collected and snap frozen. All muscles were stored at -80°C until analysis. The mouse was then killed by cardiac excision.

2.4 Taurine content analysis

The contralateral TA was freeze dried and powdered. Approximately two milligrams was powdered in a mortar and pestle and added to 20 % (wt: vol) sulfosalicyclic acid solution (Sigma Aldrich), vortexed and placed on ice. Samples were then centrifuged at 12,000 g 2oC for 2 min. Supernatant (125 μl) was collected, added to 900 μl of 0.4 M borate buffer and placed on ice before centrifuged at 12,000 g 2oC for a further 2 min. Supernatant (850 μl) was removed and stored at -80°C for HPLC analysis, which was carried out by ACS Laboratories (Kensington, Australia). Samples were analysed along with a series of standards (r2 = 0.99) and peak area was used to express taurine concentrations as μmol.g-1.

2.5 Muscle Histology

Transverse cryosections (8 μm) were cut from the midpoint of the TA and mounted on positively charged microscope slides (Lomb Scientific). Slides were stained with haematoxylin and eosin (H&E) and images were taken using a Motic BA310 microscope mounted with a Moticam 5 camera running Motic image plus 2.0 software (Motic, China). A semi-quantitative approach was taken on the contracted limb of four animals from each group, determining non-contractile area (i.e. area containing no contractile machinery including but not limited to necrotic myofibers, fragmented sarcoplasm, connective tissue and inflammatory cells) as a percentage of the total area of the section, total number of fibers per section and the number of fibers containing central nuclei. All analysis was performed blind by four parties and performed interactively, both manually and using Image J software.

2.6 Western Blotting

Frozen TA cryosections were cut from the midpoint (~30 x 10 µm 28 d animals, ~20 x 10 µm 70 d animals) and immediately placed into 1X SDS solution (3X SDS solution (0.125 M Tris-HCI, 10% glycerol, 4% SDS, 4 M urea, 10% mercaptoethanol and 0.001% bromophenol blue, pH 6.8) diluted 2:1 with 1× Tris.Cl (pH 6.8), 5mM EGTA). Samples incubated for at least 30 min at room temperature, vortexed at five minute intervals and stored at -80°C until analysed. Aliquots of each TA sample were pooled together and used to create a calibration curve that was run on every gel, allowing comparisons of whole muscle homogenates across gels23,24. Total protein from each sample was initially separated on 4-15% gradient Criterion TGX Stain Free gels (BioRad, Hercules, CA) and following UV activation using a Stain Free Imager (BioRad), the densities of the total lanes was obtained (Image lab software v 5.2, BioRad) and used to ensure equal loading for subsequent western blotting.

Quantitative western blotting was performed to determine the protein abundance of actin, myosin heavy chain 1 (MHC1), ryanodine receptor (RyR), sarco/endoplasmic reticulum Ca2+ ATPase pump 1 (SERCA1), dihydropyridine receptor (DHPR), Calsequestrin 1 (CSQ1) and 2 (CSQ2), dystrophin, utrophin, myogenin and taurine transporter (TauT). The western blotting protocol was similar to that described previously25,26,27. Briefly, an equal amount of protein from TA samples as well as a four to five point calibration curve was separated on 4-15% gradient Criterion TGX Stain Free gels (BioRad, Hercules, CA). All samples from WT, mdx and mdx tau mice were run in at least duplicate for each protein (except MHC1) and averaged across gels. Prior to transfer gels were imaged with a Stain Free Imager (BioRad) for total protein which was quantified for each sample (Image lab software v 5.2, BioRad). Following this, using a wet transfer protocol, protein was transferred onto a nitrocellulose membrane at 100V for 30 min. Following transfer the gel was imaged again and the membrane incubated in Pierce Miser solution (Pierce, Rockford, IL) for ~10 min and then blocked in 5% skim milk powder in 1% Tris-buffered, saline-Tween (TBST) for ~2 h at room temperature. Following blocking membranes were incubated in primary antibodies overnight at 4°C and 2 h at room temperature. All antibodies were diluted in 1% bovine serum albumin (BSA) in phosphate buffered saline (PBS) with 0.025% Tween (PBST).

Antibody details and dilutions are as follows: rabbit polyclonal monoclonal anti-Actin (Batch A 2066, SIGMA, 1:300); mouse monoclonal anti-MHC1 (A4.840, Developmental Studies Hybridoma Bank (DSHB), IgM, 1:200), mouse monoclonal anti-RyR1 (34C, DSHB, 1:300), mouse monoclonal anti-SERCA1 (CaF2-5D2, DSHB, 1:1,000), mouse monoclonal anti-DHPR (IIID5E1, DSHB, 1:400), mouse monoclonal anti-CSQ1 (ab2824, Abcam, 1:2000), rabbit polyclonal anti-CSQ2, (ab3516, Abcam, 1:1,000), rabbit polyclonal anti-Taurine Transporter (TauT, Professor David Pow, RMIT University, 1:10,000), mouse monoclonal anti-Utrophin (Mancho3 clone 8A4, DSHB, 1:200), mouse monoclonal anti-Dystrophin (MANDYS1 clone 3B7, DSHB, 1:500) and mouse monoclonal anti-Myogenin (F5D, DSHB, 1:250). The gel of protein extracts from mdx muscle was probed for dystrophin to confirm the absence of this protein.

After washing, membranes were incubated with a secondary antibody (goat anti-mouse IgG or IgM, goat anti-rabbit IgG, HRP conjugated, 1:60,000) and rinsed in TBST. Bands were visualized using West Femto chemiluminescent substrate (ThermoScientific, IL, USA) with images taken and densitometry performed using Image Lab software (BioRad). The positions of molecular mass markers were captured under white light before chemiluminescent imaging, without moving the membrane. Total protein and specific protein densities were each expressed relative to their respective calibration curves and subsequently each protein was normalised to the total protein content (25).

2.7 Statistics

All data are presented as mean + standard deviation (SD) unless stated otherwise. Comparisons between WT and mdx, mdx and mdx tau groups were performed using a 1-way ANOVA of variance with individual stimulation frequencies in the force-frequency analysis, and individual time points in the fatigue and recovery analyses, with Sidak’s post-hoc analyses. All statistical analysis was performed using GraphPad Prism v 6. Significance was set at p<0.05.


3.1 Effect of taurine supplementation on body mass, muscle mass, TauT protein content and taurine content

Taurine supplementation (in utero and in the drinking water) had no effect on the body mass of 28 d or 70 d WT, mdx and mdx tau mice (Table 1). The TA muscle mass was similar in 70 d mdx and mdx tau mice, and was 23% greater in the mdx mice than the WT mice (p<0.05, Table 1). At 28 d no difference in TA muscle mass was observed between WT, mdx and mdx tau groups (Table 1).

Table 1

Table 1: Body mass, tibialis anterior (TA) mass, morphological, twitch and tetanic contractile properties of TA muscles from 28 and 70 d WT, mdx and mdx taurine supplemented (mdx tau) mice.

Optimal length (Lo), cross sectional area (CSA), peak twitch force (Pt), time to peak tension (TTP) and half relaxation time (½RT). n = number of mice. Values are mean (SD); One way ANOVA with Sidak’s post-hoc analyses between age groups. Symbols for significant differences (p < 0.05) are: *significantly different from WT group, # significantly different than mdx group.

To determine the content of taurine transporter protein (TauT), over a range of loading amounts, the density of a known amount of total protein and TauT (indicated in Fig. 1A and 1B) was plotted as a calibration curve and individual samples were expressed relative to this calibration curve. As can be seen, the density of the TauT was low, however the use of the calibration curve meant that such amounts could be determined. There was no difference in the TauT abundance in the TA muscle across age or phenotype (Fig. 1C).

Muscle taurine content was measured in the TA muscle of the contralateral, non-stimulated limb of mice from each group. In the 28 d group mdx tau supplemented mice had 25% greater muscle taurine content (37 ±5 μmol.g-1) than non-supplemented mdx mice (28 ±2 μmol.g-1), and mdx mice taurine content was similar to WT mice (30 ±4 μmol.g-1, Fig 1D). Conversely at 70 d taurine supplementation proved ineffective, with mdx tau mice (39 ±7 μmol.g-1) having 22% less intramuscular taurine than the non-supplemented 70 d mdx group (47 ±6 μmol.g-1, p<0.05), and the mdx mice had similarly higher taurine levels than the 70 d WT mice (37 ±3 μmol.g-1). A significant age-specific increase in taurine content was observed with non-supplemented mdx mice increasing by 67% from 28 to 70 d (p<0.05). WT and mdx tau mice had no increase in taurine content with age (Fig. 1D).

Figure 1

Fig. 1: Taurine transporter protein and taurine content in tibialis anterior muscles from 28 d and 70 d wild-type, mdx and mdx taurine supplemented mice.

A. Western blot of taurine transporter protein (TauT, top) and Stain Free gel showing total protein (bottom) from tibialis anterior (TA) 28 and 70 d wild-type (WT), mdx and mdx taurine supplemented (mdx tau) mice. The four point calibration curve is indicated. Two different molecular weight markers are shown between the samples and the calibration curve, with the protein sizes indicated. B. Densities of total protein (left) and TauT protein (right) plotted against the calibration curve volumes loaded. The linear regression, y = mx + c is shown, along with correlation coefficient (r2). C. The TauT protein abundance is shown for 28 d (black bars) and 70 d (white bars) WT, mdx and mdx tau mice. D. TA taurine content represented as in C. Lines above specific bars indicate significant difference between groups (p<0.05). One way ANOVA with Sidak’s post-hoc analyses. Data presented as means + SD with n indicated in respective bars.

3.2 Effect of taurine supplementation on twitch and tetanic contractile properties

Figure 2A and B show the mean responses for in situ maximum and relative contractile force, respectively, for all groups. Table 1 shows the mean values for in situ twitch and tetanic TA muscle characteristics. While both maximum and specific forces in 28 d mdx mice were significantly weaker than the WT group, taurine supplementation ameliorated this weakness by approximately 60% when relative to TA muscle mass compared to the non-supplemented mdx mice (Fig. 2A, B). Whilst maximum force production was not significantly different between 70 d groups (Fig. 2A), when normalised to TA muscle mass, mdx mice produced a similar specific force to mdx tau mice, but ~27% less force than the WT mice (Fig. 2B). At 70 d taurine proved ineffective at increasing specific force production despite increasing muscle CSA by 9% (Fig. 2A, Table 1). Compared with 28 d WT, mdx mice had ~50% reduced peak twitch forces (Pt), but this was attenuated in 28 d mdx tau mice (Table 1). No differences were observed in optimum length (Lo), cross section area (CSA), time to peak tension (TTP) and half-relaxation time (½ RT) between all 28 d groups (Table 1). In 70 d animals, no differences were observed in Pt, TTP or ½ RT, but compared with mdx mice, mdx tau mice had ~9% smaller Lo (Table 1).

Figure 2

Fig. 2: In situ force production in tibialis anterior muscles from 28 d and 70 d wild-type, mdx and mdx taurine supplemented mice.

In situ maximum force (A) and specific force (B) productions in 28 d (solid symbols) and 70 d (hollow symbols) wild-type (WT), mdx and mdx taurine supplemented (mdx tau) mice. Lines above specific bars indicate significant differences between groups (p < 0.05). One way ANOVA with Sidak’s post-hoc analyses. Data presented as means + SD with n indicated.

3.3 Force Frequency Relationship

Despite taurine mice producing greater absolute forces (Fig. 2A), taurine had no effect on relative muscle force production at both low and high stimulation frequencies (p>0.05, Fig. 3). At 10 and 20 Hz, the TA muscle from mdx mice were more sensitive to the stimuli than WT animals, and similar to mdx tau mice, as indicated by producing a greater percentage of maximum force at these frequencies (Fig. 3). This trend persisted throughout the frequency range of 30 – 80Hz (Fig. 3). There was no significant difference in the force frequency curve relationship between 70 d groups (p>0.05, Fig. 3). Generally 28 d groups were more sensitive (i.e. produced a greater percentage of maximum force at a given frequency) across the lower physiological frequencies than 70 d groups. There was a significant age effect between mdx mice at 10, 20, 30, 40, 150 and 200Hz with the 28 d mdx mice more sensitive throughout (Fig. 3). Similarly 28 d mdx tau mice were more sensitive at 10, 20, 30, 40 and 50Hz when compared to the 70 d mdx tau group. 28 d WT mice were more linear throughout the frequency range than the 70 d WT.

Figure 3

Fig. 3: Force frequency relationship in 28 d and 70 d wild-type (WT), mdx and mdx taurine (mdx tau) mice.

Data are presented as means, error bars have been removed for clarity (28 d WT n= 9, 28 d mdx n=14, 28 d mdx tau n=10, 70 d WT n=5, 70 d mdx n=9, 70 d mdx tau n=8). One way ANOVA with Sidak’s post-hoc analyses within relevant groups. Symbols for significant differences (p < 0.05) are: # 28 d WT vs. 28 d mdx, ^ 28 d mdx vs. 70 d mdx, * 28 d mdx tau vs. 70 d mdx tau.

3.4 Fatigue Recovery

In response to a low stimulation fatigue protocol, the 28 d WT mice experienced an approximately 40% reduction in force production after 180 s of in situ stimulation (Fig. 4-left). In response to the same protocol 28 d mdx and mdx tau mice fatigued significantly less than the WT, with an ~20% reduction in force production (Fig. 4-left). Following 180 s recovery, 28 d WT and mdx mice produced 120% of their pre-fatigue force and mdx tau mice produced 113%. These elevated forces persisted throughout all recovery intervals (Fig. 4-right). At 70 d there was no difference in the rate or degree of fatigue between groups, with WT, mdx and mdx tau groups producing 53%, 46% and 40% of their initial force, respectively (Fig. 4-left). There was no difference in force production at any recovery interval after the 180 s fatigue protocol in 70 d mice (Fig. 4-right).

Figure 4

Fig. 4: 60 Hz fatigue and recovery in 28 d and 70 d wild-type (WT), mdx and mdx taurine (mdx tau) mice.

Data presented as means, error bars have been removed for clarity (28 d WT n= 9, 28 d mdx =14, 28 d mdx tau=10, 70 d WT=5, 70 d mdx=9, 70 d mdx tau n=8). One way ANOVA with Sidak’s post-hoc analyses within relevant groups. Symbols for significant differences (p<0.05) are: # 28 d WT vs. 28 d mdx, $ 28 d WT vs. 70 d WT, ^ 28 d mdx vs. 70 d mdx, * 28 d mdx tau vs. 70 d mdx tau.

3.5 Histopathology

To investigate the effect of age and taurine supplementation on muscle architecture the contracted TA was assessed in 28 and 70 d WT, mdx and mdx tau mice for markers of histopathology. Figure 5A shows representative transverse sections of each group. The area within each section that was comprised of non-contractile tissue (NCT) was quantified and expressed as a percentage (%) of total area (Fig. 5B). Total discernible muscle fibers were counted and the proportion of those that were centrally nucleated (CNF) was quantified (Fig. 5C). Approximately 22% of the TA cross sections from 28 d mdx mice was composed of necrotic and non-contractile tissue, with 55% of fibers being centrally nucleated and highly variable in diameter (Fig. 5). 28 d mdx tau mice had a marked reduction in histopathological features compared to the mdx mice, with ~50% reduction in NCT (Fig. 5B), a reduction in CNF (37%) and comparatively uniform muscle fiber diameter (Fig. 5C). Conversely, taurine had no effect at reducing the amount of NCT or CNF in 70 d mdx mice (Fig.s 5B, 5C), with both mdx groups visually exhibiting relatively uniform muscle fiber diameters reminiscent of the WT (Fig. 5A mdx tau 28 d and 70 d). Compared to the 28 d mdx mouse, there was both a natural 74% reduction in NCT and a marked visual improvement in fiber health in the 70 d mdx mouse. This age specific improvement was not evident in the WT or as pronounced in mdx tau groups.

Figure 5

Fig. 5: Histological characteristics of 28 d and 70 d wild-type, mdx and mdx taurine mice.

Representative haemotoxylin and eosin stained transverse sections of tibialis anterior muscles from 28 and 70 d WT, mdx and mdx tau mice shown at 400X magnification. Nuclei are stained dark, cytoplasm pink/orange. Fiber outlines are evident, non-contractile tissue (NCT) are indicated, and centrally nucleated fibers (CNF), scale bars = 100 µm. B. Quantification of the percentage area of NCT (B) and CNF (C) for 28 d (black bars) and 70 d (white bars) WT, mdx and mdx tau mice. Lines above specific bars indicate significant difference (p<0.05), One way ANOVA with Sidak’s post-hoc analyses. Data presented as means + SD, n = 4 all groups.

3.6 Effect of taurine supplementation on abundance of Ca2+ handling proteins

To investigate if the improvement in force with taurine supplementation could be attributed to altered Ca2+ handling, or the presence of an age specific effect, we measured the abundance of a number of relevant proteins, RyR1, DHPR, SERCA1, CSQ1, CSQ2, MHC1 and actin using quantitative western blotting (Table 2 and Fig.s 6 and 7). Taurine supplementation had no effect on the abundance of these proteins in either 28 or 70 d mdx mice, but there was an age specific affect in mdx and mdx tau mice, with an increase in the abundance of SERCA1 (51%) and a decrease in CSQ2 (65%) in 28 d mice and a 54% increase in CSQ1 in the mdx tau mice (Fig. 7).

Figure 6

Fig. 6: Representative blots for data shown in Table 2.

Shown for each panel is the myosin from the Stain Free gel, indicative of total protein (top) and the representative Western blot protein (bottom) for MHC1 (A, black line indicates non-contiguous lanes from the same gel), actin (B), DHPR (C) and RyR1 (D) in 28 d and 70 d WT, mdx and mdx tau mice.

Table 2

Table 2: Abundance of proteins important for excitation-contraction coupling in 28 d and 70 d wild-type (WT), mdx and mdx taurine (mdx tau) mice.

Each protein is expressed relative to density of total protein determined from the Stain Free gel and then relative to the average of 28 d WT mice. One way ANOVA with Sidak’s post-hoc analyses between relevant groups. Significance at p<0.05, data presented as mean (SD) (no significant differences observed). Representative Blots shown in Figure 6.

Figure 7

Fig. 7: SERCA1 and calsequestrin in 28 and 70 d WT, mdx and mdx tau mice.

Shown for each panel is the myosin from the Stain Free gel, indicative of total protein (top) and the representative Western blot protein (middle) and quantification of the abundance of SERCA1 (A), CSQ1 (B) and CSQ2 (C) in TA muscle from 28 d (black bars) and 70 d (white bars) WT, mdx and mdx tau mice expressed relative to the 28 d WT. One way ANOVA with Sidak’s post-hoc analyses between relevant groups. Data presented as means + SD with n indicated in respective bars. Lines connecting different bars indicate significance at p<0.05.

3.7 Effect of taurine supplementation on the abundance of pathologically relevant proteins

Dystrophin was absent in all mdx groups and present in the WT confirming the phenotype (Fig. 8A). Taurine supplementation had no effect on the abundance of utrophin or myogenin at 28 d (Fig.s 8B, 8C). Utrophin was 180% greater in 28 d mdx mice compared to the 70 d mdx mice (Fig. 8B). Interestingly, myogenin abundance in 70 d mdx tau mice was 450% and 230% greater than 28 d mdx tau and 70 d mdx mice, respectively (Fig. 8C).

Figure 8

Fig. 8: Dystrophin, utrophin and myogenin in 28 and 70 d WT, mdx and mdx tau mice.

Shown for each panel is the myosin from the Stain Free gel, indicative of total protein (top) and the representative Western blot protein (middle) and quantification of the abundance of dystrophin (A), utrophin (B) and myogenin (C) in TA muscle from 28 d (black bars) and 70 d (white bars) WT, mdx and mdx tau mice expressed relative to the 28 d WT. One way ANOVA with Sidak’s post-hoc analyses between relevant groups. Data presented as means + SD with n indicated in respective bars. Lines connecting different bars indicate significance at p<0.05.


The major aim of this study was to compare the efficacy of taurine supplementation given prior to conception at ameliorating dystrophic symptoms in the 28 and 70 d mdx mouse. Here we have demonstrated taurine to be beneficial during the acute damage phase at 28 d, but not at 70 d where the pathology of the mdx mouse has become stable. These findings support the potential of taurine to act in a protective role in the treatment of DMD, and highlight the importance of age as a key consideration when utilising the mdx mouse when screening for therapeutic supplements and therapies.

Oral taurine supplementation given to mothers, then to weaned pups, successfully elevated taurine content of the TA muscle in 28 d but not 70 d mdx mice. Interestingly, 28 d mdx mice had endogenous levels of taurine similar to WT mice, and 70 d mdx mice had significantly greater muscle taurine content than both WT and mdx tau mice (Fig. 1D). Whilst not strictly comparable due to different methods of analysis, McIntosh et al28 previously found that the intramuscular taurine content of 3-6 week old mdx mice was lower than WT mice, although in that study the concentration of taurine was ~10-fold less than the values obtained here (2.17 and 2.61 μmol.g-1, in WT and mdx mice, respectively), suggesting that the total pool of taurine was not measured in that study. In line with the current work, intramuscular taurine content was similar in quadriceps muscle from 22 d mdx and WT mice13 and 42 d mdx and WT mice (~24 μmol.g-1)17, although decreased in 28 d mdx compared with WT mice17. Whilst we report an age-related increase in TA intramuscular taurine content in mdx mice, in our hands there was no increase in muscle taurine content in 70 d mdx tau mice. When supplementation began at 18 d, 24 days of taurine supplementation in mdx mice increased TA taurine content16, suggesting that the lack of taurine increase we observed was due to events in the last four weeks. That taurine did not improve the contractility of mdx muscle at 70 d suggests that the beneficial effects of taurine are not independent of the pathology. Indeed, the lower taurine content in 70 d mdx tau mice suggests there must be a decrease in taurine uptake in those mice. Cozzoli et al29 reported elevated muscle taurine content in 8-12 week old taurine supplemented mdx mice compared to mdx controls, although given the 5 week age differential of mice used in that study, it is difficult to interpret further. A further difference from our current study is that those animals had undergone damaging exercise protocols to increase the severity of the mdx phenotype, a commonly used intervention, although it is not known if that intervention may affect the intramuscular taurine content, for example, by the muscle compensating for any taurine loss due to muscle damage, which makes direct comparisons not possible. The similar muscle taurine content in 70 d mdx tau mice which we observed could be due to the relative muscle health of mdx tau mice already seen at 28 d (see discussion on histopathology) possibly carrying through to adulthood (Fig. 1B, Fig. 5A, see mdx tau 28 d).

In this study we found no difference in the abundance of the TauT protein between WT, mdx and mdx tau mice at either 28 d or 70 d. Given the change in intramuscular taurine content, this finding suggests that the activity of the protein is regulated via post-translational modifications. Terrill et al17 similarly identified that TauT protein content in quadriceps muscle from non-supplemented mdx mice did not change with age, however TauT protein content was found to be consistently lower than 18, 28 and 42 day age matched WT mice. It should be noted that differences in sample preparation may explain some of the disparity between the two studies and so direct comparison may not be valid. De Luca et al30 found an increase in plasma taurine content in 6-8 month old mdx mice suggesting issues with the function of the TauT in uptake or retention in the muscle. Given we have only measured the TauT protein abundance we are not able to comment on any potential difference in the activity of the transporter.

We have uniquely measured in situ force production in 28 d mdx mice with and without taurine supplementation. Investigating contractile characteristics in situ affords many translational benefits. Having nerve and blood flow intact, maintaining normal temperatures and incorporating nerve stimulation provide an ideal situation for assessing isometric force production in a physiologically relevant setting (See SOP in methods). When looking at contractile characteristics in situ, compared with mdx mice there was a significant increase in peak twitch (Pt) and a mild, yet not significant, increase in maximum force in 28 d mdx tau mice, with no such differences in 70 d mdx and mdx tau mice (Table 1 and Fig. 2A). When expressed as specific force which takes into account the ~23% greater muscle mass, the TA muscles from 28 d mdx mice were weaker than both the WT and mdx tau mice (Table 1, Fig. 2B). In 70 d mice, there was no difference in maximum force between the groups, although the mdx mice had significantly reduced specific force than the WT, but not different from the mdx tau mice (Fig. 2B). A reduction in force production despite an increase in muscle mass would classically be considered as pseudohypertrophy, whereby a large proportion of the muscle is replaced by fat and connective tissue. Rather, visual analysis of muscle from 70 d mdx and mdx tau mice reveals a low occurrence of fibrosis and myofibers of increased size representative of true hypertrophy, and not pseudohypertrophy (Fig 5A – mdx 70 d). Investigating strength relative to muscle size is important as, similar to boys with DMD, regenerating mdx mice can undergo a compensatory hypertrophy of skeletal muscle to circumvent weakness31,32. In absolute terms, muscle from mdx mice has previously been found to be comparative in strength to WT mice, although weaker when expressed as specific force33. Different methodologies make it difficult to draw direct comparisons, however previous studies investigating taurine supplementation in mdx mice aged 6-12 weeks have similarly reported it to be efficacious at enhancing grip strength in vivo and in isolated muscle preparation ex vivo16,20,29.

At 28 d mdx mice produced a greater proportion of their maximum force at lower stimulation frequencies (10 and 20 Hz) compared to the WT, with there being no difference between mdx and mdx tau mice (Fig. 3). There was no phenotype differences apparent in 70 d mice, however across most frequencies measured, the force produced expressed as a percentage of maximum force was typically greater in the 28 d and 70 d mdx mice compared with the age-matched mdx tau mice (Fig. 3). Our finding in the younger mice is consistent with prior studies finding mdx mice to having a more negative mechanical threshold (MT) for excitation-contraction coupling (E-C coupling)34. Briefly, E-C coupling takes place when a depolarisation event activates the voltage sensing dihydropyridine receptor (DHPR) which causes Ca2+ release from the sarcoplasmic reticulum (SR) via pressing on the ryanodine receptor (RyR). Ca2+ subsequently activates the contractile apparatus, stopping when sufficient Ca2+ is sequestered back into the SR via the Ca2+ ATPase pump (SERCA) where the Ca2+ is bound to calsequestrin (CSQ)35. There is evidence that mdx mice conduct action potentials normally but subsequent SR Ca2+ release is moderately reduced36,37,38. This reduction may be compensatory, as cytosolic Ca2+ content in mdx mice resulting from membrane tears and/or ion channel dysfunction is elevated, thus reducing the amount of Ca2+ release required for contraction and resulting in a decrease in the MT30. While speculative this allows dystrophic muscle to contract more easily, the deleterious effects associated with disrupted Ca2+ homeostasis, such as activation of Ca2+ dependent proteases, make ameliorating the MT a desirable outcome when assessing DMD treatment options. In adult mdx mice, taurine has previously been found to ameliorate the MT in vitro, both when applied directly to whole EDL preparations and in EDL muscle fibers from taurine supplemented mdx mice that had undergone damaging exercise to induce the severity of the phenotype20,30. In our in situ model, comparing the percentage of maximum force at a given frequency along with similar abundance of proteins involved in E-C coupling and Ca2+ regulation, we found that in both 28 d and 70 d mdx mice, taurine was ineffective at influencing Ca2+ homeostasis.

Despite the increase in maximum contractile force in the 28 d mice, taurine had no effect on TTP or ½RT (Table 1) which can be crude indicators of the rates of Ca2+ release and re-uptake by the ryanodine receptor (RyR, or calcium release channel) and SERCA pump, respectively. Using western blotting, we report no change in the abundances of RyR (Table 2) and SERCA (Fig. 7) proteins with age or phenotype, although the protein contents were highly variable. The similar abundances of RyR1 and SERCA1 further supports the similar levels of Ca2+ homeostasis discussed above. Additionally, our finding in the 70 d mice is consistent with a prior study in slightly younger, 8 week old mdx mice where no differences in these proteins were observed39. However, while content levels are consistent, we cannot preclude any impact of taurine on the activities, or efficiencies, of these proteins. Indeed SERCA activity is suppressed in mdx mice as well as double knock-out mice, which are deficient in both dystrophin and utrophin40, and it is possible that taurine could improve SERCA activity and warrants further examination in these models. The abundance of CSQ2, a high capacity Ca2+ binding protein located in the SR (typically associated with slow twitch muscle), was ~3-fold greater in 28 d mdx compared with WT mice, and was similar to the CSQ2 content in mdx tau mice (Fig. 7). By 70 d of age, the amount of CSQ2 was no longer different between phenotypes (Fig. 7). The increased CSQ2 abundance in 28 d mdx mice may point towards a shift in the muscle isoform, however we found no differences in the abundance of the slow myosin isoform (MHC1, Table 2) or in the abundance of CSQ1, the CSQ isoform dominating in fast-twitch muscle (Fig. 7), suggesting this is not the case. Murphy et al25 suggested CSQ2 may have a role in reducing the amount of free Ca2+ within the SR, which would prevent SR leak from further exacerbating the high cytosolic Ca2+ already associated with dystrophy. Previously the abundance of total CSQ (i.e. CSQ1 and CSQ2) was shown to be reduced in 9 week old mdx mice and it was suggested that this contributes to a reduced Ca2+ buffering capacity41. We found no such reduction in the abundance of CSQ isoforms in either 28 d or 70 d mice, however it is difficult to compare the findings from the two studies because in the previous work total muscle extract preparation seemingly involved removal of fractions through filtration or centrifugation, which even at low centrifugal force has been shown to result in loss of up to 60% of the total CSQ in cardiac tissue26 and (ii) the CSQ isoforms, CSQ1 and CSQ2, were not distinguished from each other as they were here.

In response to the 60 Hz in situ fatigue/recovery protocol 28 d mdx mice were more resistant to fatigue during 180 s of the intermittent low frequency stimulation than WT mice and this was not different in mdx tau mice (Fig. 4). There was no difference observed between the 70 d groups. Generally mdx mice are reported to be highly susceptible to fatigue although this is dependent on mouse age and the nature of the fatigue protocol31,33. Sacco et al 42 repeatedly stimulated the TA of adult female mdx mice at 40 Hz and similar to the current study, reported that mdx mice were more resilient to fatigue than the WT mice. Taken together, our findings suggest that the weaker mdx mice, as measured by a decreased specific force (Fig. 2B), require less ATP and can thus spare energy for further contractions. Due to the smaller muscle size in 28 d mice, it would be expected that a larger percentage of the muscle is recruited during a 60 Hz stimulation compared with the larger 70 d animals, which would be expected to result in the muscle fatiguing more readily, although this was not observed. Interestingly, all 28 d animals, but not 70 d animals, experienced an increase in maximal force production following the fatigue protocol (Fig. 4-right). This potentiation is due to increased phosphorylation of the myosin light chains following repeated stimulation, subsequently making the contractile filaments more sensitive to Ca2+ creating more, as well as greater, cross bridging43,44. Given that the supra maximal force post fatigue was also observed in the 28 d WT mice excludes a pathological based explanation for the potentiation, leaving large muscle motor unit recruitment due to small muscle size as a likely explanation. Previously a higher susceptibility to fatigue was reported in mdx mice, but this was assessed in the diaphragm which, unlike hindlimb muscles of these animals that regenerate into adulthood, continues to degenerate throughout the life the mdx45. Grip strength of 3-24 week old mdx mice fatigued more rapidly than WT mice46 however when considering the avoidance behaviour mdx mice exhibit towards exercise9,47 it is unclear if this data is a true physiological representation of muscular endurance. The fatiguing properties of EDL muscle were reported to be greater at physiological temperature (35oC compared with 20oC)48. Using an in situ protocol we have addressed physiological temperature, kept nerve and blood flow intact and circumvented avoidance behaviour to get a true indication of muscular endurance capacity and at this frequency we found no difference in the fatigability of mdx mice at 70 d of age. Given that fatigue and recovery were not compromised in 70 d mdx mice it was perhaps not surprising to see no difference in mdx tau mice. This is in contrast to studies in rats, where taurine supplementation improved fatigability21,49.

There appears to be a link between increased strength and improved histological profile and visible health in 28 d mdx tau compared with mdx mice, with similar improvements seen with age (Fig. 5). Non-supplemented 28 d mdx mice displayed classic markers of dystrophic damage with large areas of non-contractile and necrotic tissue with a prevalence of recently regenerating, centrally nucleated myofibers. Speculatively, the force deficit seen in the mdx mouse may be due to the loss of healthy, functional contractile tissue seen here. However this may also be due to the high proportion of immature myofibers seen at 28 d, which are a result of the acute onset of myofiber necrosis the model experiences at approximately 21 d9. In contrast there is little evidence of active necrosis in 28 d mdx tau mice, which have approximately half the amount of non-contractile tissue seen in the mdx group and subsequently produced approximately twice the amount of force. There was also a decrease in the number of centrally nucleated fibers seen in mdx tau mice when compared to mdx mice. A similar result was found by Terrill and colleagues13, when supplementing mdx mice with taurine from 14 days of age, finding a reduction in myofiber necrosis and inflammation, also finding there was little to no marked presence of centrally nucleated myofibers in 22 d juvenile mdx mice. Previously mdx mice with higher taurine content have also been found to have the most effective muscle regeneration50. In the current study it is unclear why the large, seemingly mature myofibers in 28 d mdx tau mice are centrally nucleated, despite no evidence of active necrosis or other recent artefacts of muscle damage. It may be explained by some developmental alteration as a result of in utero taurine supplementation or that necrosis occurred at a time point early enough to allow the development of a mature myofiber or that taurine is facilitating a more rapid regenerative response.

In muscle from 70 d mdx and mdx tau mice, there are large, recently regenerated myofibers and it is likely that the central nuclei are an artefact of prior, rather than current damage. Those mice also display a more uniform muscle fiber diameter and a comparable amount of non-contractile tissue to the WT mice, providing an explanation for why, in the most part, they are capable of conferring more mechanical strength than the smaller, recently regenerated myofibers of the 28 d mdx mice (Fig. 5A, compare white space and clumped nuclei in mdx 28 d, and relative fiber size in mdx tau 28 d). Similar to contractile strength, the improved histology seen between mdx and mdx tau mice at 28 d was not evident in mdx and mdx tau mice at 70 d. This was not surprising, as it is well-documented that 10-14 week old mdx mice have little dystrophic phenotype due to the successful regenerative period. The inclusion of this age group in the current study, was to highlight that these mice, unless chronically exercised to aggravate the pathology, as widely used9, are not a good dystrophic model, whilst the 28 d mdx mouse presents an appropriate model.

As a measure of myofiber health, we also examined the abundance of myogenin protein, a myogenic regulatory factor involved in skeletal muscle health, development and regeneration. In 28 d mice, the abundance of myogenin was variable and not significantly different between groups of mice; however in 70 d mdx tau mice there was a ~3-fold increase in myogenin protein content compared with mdx mice. Previously it was reported that myogenin was dispensable for skeletal muscle health in adult mdx mice although it appeared to be elevated in response to muscle degeneration in juvenile mdx mice51. Certainly, at the mRNA level, compared with WT mice, myogenin was elevated in 21 d mdx mice and seemingly sustained as mice aged to over 12 months, although the latter finding was reported from a total of two animals52. Whilst our findings of increased myogenin content in mdx tau mice may indicate improved muscle health in the taurine supplemented 70 d mice, the upregulation of myogenin in 70 d mdx tau occurred despite there being no visible or quantifiable difference in the myofiber health between mdx and mdx tau mice. It is widely recognised that elevated myogenin, along with other markers such as increased acetylcholine receptor abundance are markers of denervation that also occur in ageing muscles53. Since denervation of the neuromuscular junctions has been reported in muscle from mdx mice the elevated myogenin in mdx tau animals (at 70 d) may reflect this denervation, rather than being due to a disturbed aspect of myogenesis54,55. Further, the findings might indicate that myogenin levels may not be down regulated in the regenerated muscle from older mdx mice. Given that intramuscular myogenin and taurine contents have independently been associated with healthy muscle development and function, it appears that any synergistic benefit is below the measurable limits of the muscle histopathology and force production.


The mdx mouse is an invaluable tool to study both the mechanisms of dystrophic damage, and assess potential therapeutic supplements and therapies to remedy the severity of DMD. The age dependent disease severity of the mdx mouse affords the unique opportunity for both the assessment of therapies to prevent or delay dystrophic symptoms (21 – 28 d), as well as treat the established disease state (70 d). This study demonstrates prenatal taurine supplementation to be efficacious at ameliorating muscle weakness and improving histological characteristics in the mdx mouse model of DMD during the acute stage myofibre necrosis seen at 28 d, but not at 70 d where the background pathology is initially mild. The complexity of DMD makes identifying supplements and therapies to target specific pathways difficult. Whilst not able to completely ameliorate the many pathological consequences of DMD, the beneficial effect of taurine seemingly lies in its ability to remedy many of the downstream pathways such as Ca2+ handling, anti-inflammation and anti-oxidation simultaneously, cumulatively reducing the severity of the pathology. Considering these findings and that taurine is a cheap, readily accessible and side effect free dietary supplement, we envisage taurine could be administered to pregnant mothers in a protective capacity, reminiscent of folate in the prevention of spinal bifida.

Corresponding Author

Dr. Robyn M. Murphy

Department of Biochemistry and Genetics, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, VIC 3086, Australia.


Competing Interests

There are no competing interests associated with this study.

Data Availability

The data sets associated with this project are available on figshare:

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Three Novel Immune-deficient Mouse Models of Muscular Dystrophy Fri, 01 Sep 2017 12:57:42 +0000 To facilitate gene and cell therapy experiments, we created severely immune-deficient mouse models of Duchenne muscular dystrophy (DMD), limb girdle muscular dystrophy 2B (LGMD2B), and limb girdle muscular dystrophy 2D (LGMD2D) by crossing mdx4Cv, Bl/AJ, and Sgca-null mice with NRG immune-deficient mice. The resulting mdx4Cv/NRG, BlAJ/NRG, and Sgca/NRG mice demonstrated the presence of the appropriate mutant alleles at Dmd, Dysf, Sgca, Rag1, and Il2 by genotyping PCR. Absence of dystrophin, dysferlin, or α-sarcoglycan protein was confirmed by western blot and immunohistochemistry. We performed centronucleation, Evans blue dye, hydroxyproline, and treadmill assays on the disease model mice versus NRG controls to evaluate muscle histology and function. These studies demonstrated that the mdx4Cv/NRG and Sgca/NRG mice showed significant deficits in muscle structure and function in all the assays and were similar to each other. By contrast, the phenotype of the BlAJ/NRG mice was milder in each case. The results we observed parallel the phenotypes seen in patients with the corresponding disorders. These novel immune-deficient mouse models of DMD, LGMD2B, and LGMD2D will be useful for long-term gene and cell therapy studies involving transfer of foreign genes and cells.



Muscular dystrophies are genetic diseases caused by mutations in genes encoding muscle proteins, leading to progressive muscle degeneration 1. Duchenne muscular dystrophy (DMD) is the most common form of muscular dystrophy, characterized by widespread degeneration of the skeletal, respiratory, and cardiac muscles, resulting in disability and premature death. DMD is caused by mutation of the dystrophin gene (DMD), which resides on the X chromosome 1,2. Limb girdle muscular dystrophy 2D (LGMD2D) is a less common autosomal recessive form of muscular dystrophy resulting from mutation of the α-sarcoglycan gene (SGCA) on chromosome 17. The phenotype of this disease is variable, depending on the nature of the mutations involved, with severe forms being similar to DMD 1,3. Limb girdle muscular dystrophy 2B (LGMD2B) is another autosomal recessive form of muscular dystrophy resulting from mutation of the dysferlin gene (DYSF) on chromosome 2. LGMD2B leads to a later-onset, milder form of muscular dystrophy characterized by progressive degeneration of skeletal muscles and resulting disability 1,4.

Because of their monoallelic genetic basis, the addition of genes or cells that could provide the DMD, SGCA, or DYSF protein that is deficient in these disorders has the potential to be an effective treatment. Gene therapy strategies to introduce functional DMD, SGCA, or DYSF genes to diseased muscle fibers are attractive approaches currently under development5,6,7,8,9. Even so, gene therapy may have diminishing efficacy as patients progressively lose muscle fibers with time. Moreover, gene therapy strategies typically do not correct muscle stem cells 10. In these settings, cell therapy strategies designed to restore healthy muscle cells may have value. If healthy muscle progenitor cells were introduced and could fuse to existing deficient fibers, as well as form new muscle fibers, regeneration of muscle might occur 11,12. The well-established ability of satellite cells and their myoblast progeny to engraft in skeletal muscle 13,14 provides a basis for cell therapy strategies, as does the use of muscle progenitor cells derived from pluripotent stem cells and other cells 15,16,17,18,19,20,21,22,23,24,25,26,27.

Cell therapy strategies require transplantation of living cells, which may involve introduction of foreign genes into the recipient organism. Gene therapy strategies are also often assisted by inclusion of foreign marker genes, such as luciferase. To create mouse models that would be appropriate for such experiments, including the transplantation of human cells into mice, we took advantage of recently created mouse strains that are severely immune deficient, lacking B, T, and NK cells. The NRG strain background is useful for muscle studies, since it tolerates irradiation 28. Ionizing radiation is sometimes used in muscle regeneration studies to eliminate the endogenous host stem cells, known as satellite cells, thereby facilitating engraftment of donor cells 29. The development of gene and cell therapies is facilitated in the background of immune-deficient animals, by removing rejection barriers and allowing the use of foreign markers and cells, thus expanding the range of permissible experimental designs. This study provides three novel severely immune-deficient mouse muscular dystrophy models that will be useful in the development of successful gene and cell therapies. We also characterize the models to provide valuable baseline phenotypic information about them. Comparing the results between strains may elucidate further the phenotypic differences between these three forms of muscular dystrophy, although we note that since the strains are not fully inbred, differences in modifier genes may be present between them that could affect phenotype.

Materials and Methods

Ethics statement

The Stanford Administrative Panel on Laboratory Animal Care approved all procedures performed on animals in protocol number 15766, assurance number A3213-01. The Stanford Comparative Medicine program is accredited by the Assessment of Laboratory Animal Care International.

Mouse strains

NOD (NOD/ShiLtJ, 001976), NRG (NOD.Cj-Rag1tmMomIl2rγtm1Wjl/Szj, 007799) 28, mdx4Cv (B6Ros.Cg-Dmdmdx-4Cv/J, 002378) 30, and mdx/scid (B10ScSn.Cg-PrkdcscidDmdmdx/J) mice were purchased from the Jackson Laboratory (Bar Harbor, ME). Bl/AJ mice (B6.A-Dysfprmd/GeneJ, 012767) 6 were provided by the Jackson Laboratory from a stock maintained by the Jain Foundation. SGCA-null mice 3 were a kind gift from Kevin Campbell.

PCR analysis of genotypes

Genomic DNA was extracted from mouse ear punch using Wizard Genomic DNA Purification Kit according to manufacture’s instructions (Promega, Cat. A1120). IL2rγ and Rag1 regions were amplified from 100 ng of genomic DNA (5 min at 95°C; 30 sec at 94°C, 30 sec at 51°C (IL2rγ) and 52°C (Rag1), 30 sec at 72°C, 35 cycles; 5 min at 72°C). The primers used to detect wild-type or mutant alleles are published on The Jackson Laboratory website. The primer sequences IL2RG-Common-Fwd (5’-AAGAGATTACTTCTGGCTGTCAG-3’) and IL2RG-Wt-Rev (5’-CTCTGGGGTTTCTGGGG-3’) were used to detect IL2rγ wild-type. The primers IL2RG-Common-Fwd and IL2RG-Mut-Rev (5’-ATGCTCCAGACTGCCTTG-3’) were used to detect IL2rγ mutant. The primer sequences Rag1-wt-Fwd (5’-TCTGGACTTGCCTCCTCTGT-3’) and Rag1-Common-Rev (5’-CATTCCATCGCAAGACTCCT-3’) were used to detect Rag1 wild-type. The primers Rag1-Common-Rev and Rag1-Mut-Fwd (5’-TGGATGTGGAATGTGTGCGAG-3’) were used to detect Rag1 mutant.

The following primer sets were used to identify mutant muscle disease gene alleles: Dysfprmd (5’-TTCCTCTCTTGTCGGTCTAG-3’), (5’-CTTCACTGGGAAGTATGTCG-3’) and (5’-GCCTTGATCAGAGTAACTGTC-3’); mdx4Cv (5’-TCAAGAACAGCTGCAGAACAGGAGA-3’) and (5’-GGATTGCATCTACTGTGTGAGGACC-3’); and Sgca (5’-GCCAGAGGCCACTTGTGTAG-3’) and (5’-ACTCACCTACCACGCTCACC-3’). Expected fragment sizes are given in Fig. 1b & c, Fig. 2b & c,and Fig. 3b & c.


Muscles were harvested and snap frozen in OCT in liquid nitrogen. Serial 8 μm cryostat sections were obtained throughout the muscle. Sections were fixed in 2% PFA for 10 minutes and washed 3X in PBS with 2% triton (PBS-T) for 1-2 min. Sections were blocked in 5% donkey serum in PBS-T for 1 hour. Primary antibodies were prepared in block solution and incubated overnight at 4°C in a humid chamber. The next day, slides were washed 3 times with PBS and stained with secondary antibody for 1 hour at room temperature. Slides were washed with PBS 3 times and visualized. Antibodies used were: rabbit anti-dystrophin (15277, Abcam, Cambridge, UK), rabbit anti-dysferlin (NCL-Hamlet Novocastra, Leica), and rabbit anti-Sgca (189254, Abcam). Donkey anti-rabbit conjugated to Alexa 594 (A21207, Life Technologies) was used as secondary antibody.

Western blots

Muscle lysates were prepared with RIPA Buffer supplemented with HALT Protease Inhibitor Cocktail (Thermo Fisher Scientific 78430, Waltham, MA) according to the manufacturer’s protocol. The supernatant containing protein extract was denatured with Laemmli Sample Buffer (Bio-Rad 1610737, Hercules, CA) supplemented with 100 mM DTT. In each lane of a 10% TRIS-glycine SDS-PAGE gel (Bio-Rad), 25 μg protein extract was electrophoresed at 70 V in ice cold PAGE running buffer (0.1% SDS, 25mM Tris, 250mM glycine). Samples were transferred onto 0.45 μm PVDF membrane (Thermo Fisher) for 50 minutes at 100 mA at 4°C. Membranes were blocked in 0.2% BSA and 2% milk diluted in TBS with 0.5% Tween-20. Dystrophin, dysferlin, and Sgca detections were respectively achieved with a 1:1000 dilution of rabbit-anti-dystrophin (15277, Abcam), rabbit-anti-dysferlin (124684, Abcam), and rabbit-anti-SGCA (189254, Abcam). GAPDH was probed with a 1:5000 dilution of rabbit-anti-GAPDH (181602, Abcam) in blocking solution. Rabbit antibodies were probed with a 1:5000 dilution of goat-anti-rabbit IgG HRP secondary antibody (Thermo Fisher). Blots were developed in Clarify Western ECL Substrate according to manufacturer’s protocol (Bio-Rad) and imaged using the ChemiDoc Touch Imaging System (Bio-Rad).

Histological staining

Tibialis anterior (TA), quadriceps (Q), hamstring (H), and gastrocnemius (G) muscles from NRG, mdx4Cv/NRG, BlAJ/NRG, and Sgca/NRG mice were isolated and flash frozen in mounting media (OCT, Satura Finetek, Torrance, CA). Cryosections 12 um-thick were stained using hematoxylin and eosin (Sigma, St. Louis, MO) following the manufacturer’s specifications. All pictures were taken using a Zeiss Aptiskop microscope.

To evaluate the membrane permeability of muscle, mice were intraperitoneally injected with Evans Blue dye (EBD; Sigma; 100 ul of 1% EBD in PBS per 10 g body weight). The following day, muscles were flash-frozen and ground with a mortar and pestle. Following an overnight incubation in formamide, EBD was quantified in the supernatant by measuring light emission at 630 nm 31.

Hydroxyproline, which is a direct measure of the amount of collagen/gelatin, was used to determine the amount of fibrosis in whole muscle. Hydroxyproline was measured according the manufacturer’s protocol (Kit-555; BioVision, Milpitas, CA). Briefly, muscles were dissected and incubated in 6N HCl for 3 hours at 120°C. Small volumes of the hydrolysates were then incubated with the Chloramine T and p–dimethylaminobenzaldehyde (DMAB) reagents. The resulting product, a chromogen, was then measured at 560 nm.


To measure mouse fatigue, mice were run on a treadmill (IITC Model 800; IITC Life Science, Woodland Hills, CA) with a 10° uphill incline. An electric shock bar grid at the end of the tread delivered a mild shock to mice if they stopped running. The fatigue time was measured as the time a mouse could run before hitting the electric shock grid 5 times. During the first week, mice were acclimated 3 times (Monday, Wednesday, Friday) at 8 m/min for 3 min. The following week, mice were similarly run 3 times with the speed set at 10-18 m/min, with increments of 2 m/min. The time that a mouse hit the electric shock grid for the 5th time was recorded as the run length. For each mouse, the times of the 3 runs were averaged. Measurements were performed at the same time each day to reduce variability.

Statistical analyses

To determine statistical significance for two groups, comparisons were made using a Student’s t-test. Statistical analyses were performed using GraphPad Prism v.6 (GraphPad Software, La Jolla, CA, USA). A p-value < 0.05 was considered significant.


Construction of immune-deficient mdx4Cv, Bl/AJ, and Sgca-null mouse models

Immune-deficient mouse models are useful in cell therapy studies since they allow transplantation of cells expressing markers that would otherwise cause rejection, preventing long-term engraftment. Such markers include species-specific antigens such as those present on human cells. Immune-deficient mouse models are also valuable in gene therapy studies, enabling the transfer of genes that might otherwise be immunogenic, including human therapeutic genes and bioluminescent and fluorescent marker genes. An immune-deficient mdx4Cv/NSG mouse line was previously constructed and has been valuable 32. On the other hand, the NSG line of mice is highly susceptible to radiation-induced damage because of mutation of the Prkc gene, which is necessary for double-strand break repair. Since irradiation is a frequently used technique for pre-conditioning skeletal muscle for cell engraftment 29, we used the NRG strain, which is less sensitive to radiation, permitting more flexibility on dosage28. Lymphocytes are eliminated in NRG mice because of a mutation in the recombination-activating gene (Rag1), which is required for VDJ recombination in lymphocytes and is immune specific 28. A different immune-deficient DMD model with similar properties, featuring a non-revertable dmd mutation and Rag2 and Il2rb mutant alleles has also been reported33.

We generated three lines of dystrophic, immune-compromised mice; one that lacks dystrophin, one that lacks dysferlin, and another that lacks α-sarcoglycan. To generate mdx4Cv/NRG mice, we combined null mutations in the genes encoding rag1, interleukin 2 receptor gamma subchain (IL2rγ), and the mdx4Cv mutation in dystrophin (dmdmdx4Cv) that introduces a premature stop codon in exon 53 of the dmd gene. The dmdmdx4Cv mutation has a 10-fold lower reversion frequency than the original mdx mutation 30,34D. Details of the crosses involved are shown in Fig. 1a. The dmdmdx4Cv mutation was monitored in crosses by using PCR (Fig. 1c). To initiate breeding, C57Bl/6/mdx4Cv/4Cv females were first outcrossed with NOD males. Female progeny were outcrossed to NOD males, and this cross was repeated three times, generating backcross generations N1, N2, and N3. N3 generation NOD/dmdmdx4Cv/+ females were outcrossed twice to NRG males to obtain backcross generations N4 and N5 dmdmdx4Cv/NRG mice. The mice progressively changed in coat color from black to white through the crosses. In addition to the dystrophic dmdmdx4Cv allele, Rag1 and Il2rg mutations were monitored by PCR and maintained beginning with the N4 generation (Fig. 1b). N5 generation NRG/dmdmdx4Cv/+ mice were intercrossed to obtain dmdmdx4Cv/NRG breeding pairs homozygous for the dmdmdx4Cv, Rag1, and Il2rγ mutations. These mice were used to propagate and maintain by inbreeding the line, designated mdx4Cv/NRG.

Figure 2

Fig. 1: Characterization of mdx4Cv/NRG mouse model

(a) Breeding scheme for generation of the mdx4Cv/NRG mouse colony. (b,c) PCR analysis showing the fragments used for identification of the NRG and mdx4Cv genotypes. Sizes of diagnostic fragments are noted in basepairs (bp). (d) Immunohistochemistry for dystrophin in gastrocnemius muscle shows the absence of dystrophin in mdx4Cv/NRG compared to NRG. (e) Western blot illustrating absence of dystrophin in mdx4Cv/NRG.

To generate BlAJ/NRG and Sgca/NRG mice, a similar breeding scheme was followed, but incorporating the Dysfprmd mutation in dysferlin or the Sgca-/- mutation in α-sarcoglycan instead of mdx4Cv(Fig. 2a, 3a). The Dysfprmd mutation and Sgca-/- mutation were obtained respectively from BlA/J mice 6 and Sgca-null mice 3. Dysfprmd, Sgca, Rag1 and Il2rγ mutations were monitored in crosses by using PCR (Fig. 2b & c; Fig. 3b & c). To initiate breeding, B6.A-Dysfprmd/GeneJ (Bl/AJ) females were first outcrossed with NOD males, and subsequent crosses were similar to those described above for establishment of the mdx4Cv/NRG mouse colony, producing the BlAJ/NRG mouse line (Fig. 2a).A similar breeding scheme was used to obtain Sgca/NRG mice from Sgca-/- females (Fig. 3a).

Figure 1

Fig. 2: Characterization of BlAJ/NRG mouse model

(a) Breeding scheme for generation of the BlAJ/NRG mouse colony. (b,c) PCR analysis showing the fragments used for identification of the NRG and Bl/AJ genotypes. Sizes of diagnostic fragments are noted in basepairs (bp). (d) Immunohistochemistry for dysferlin in gastrocnemius muscle of NRG and BlAJ/NRG demonstrated the absence of dysferlin in BlAJ/NRG. (e) Western blot illustrating absence of dysferlin in BlAJ/NRG.

Figure 3

Fig. 3: Characterization of Sgca/NRG mouse model.

(a) Breeding scheme for generation of the Sgca/NRG mouse colony. (b,c) PCR analysis showing the fragments used for identification of the NRG and Sgca-null genotypes. Sizes of diagnostic fragments are noted in basepairs (bp). (d) Immunohistochemistry for a-sarcoglycan in gastrocnemius muscle of NRGand Sgca/NRG demonstrated the absence of a-sarcoglycan in Sgca/NRG. (e) Western blot illustrating absence of a-sarcoglycan in Sgca/NRG.

Muscle studies

The muscle phenotype of mdx4Cv/NRG mice was assessed in 6 month-old animals. Muscle sections from mdx4Cv/NRG and NRG mice were stained with antibodies that bound specifically to dystrophin. As shown in Fig. 1d, dystrophin was present at the sarcolemma of all fibers in NRG muscle, but was completely absent in muscle from mdx4Cv/NRG (Fig. 1d), validating the absence of this membrane-associated muscle protein in the disease model mice. Western blots further substantiated the absence of dystrophin (Fig. 1e).

To evaluate the BlAJ/NRG mice, muscle sections from 6 month-old BlAJ/NRG mice and the NRG control were stained with antibodies that bound specifically to dysferlin. We observed dysferlin staining predominantly localized to the membrane in the NRG mice (Fig. 2d), but an absence of staining in the BlAJ/NRG mice (Fig. 2d), consistent with a null phenotype for dysferlin. Western blots for dysferlin from the two samples confirmed the absence of dysferlin in the BlAJ/NRG mice (Fig. 2e). Likewise, immunohistochemistry (Fig. 3d) and western blotting (Fig 3e) confirmed the absence of Sgca protein in the Sgca/NRG mouse model.

To analyze any associated muscle pathology, hematoxylin and eosin (H&E) staining was carried out on muscle sections from the three mouse models. As seen in Fig. 4a, the NRG muscle appeared normal, while the muscles from mdx4Cv/NRG, BlAJ/NRG and Sgca/NRG mouse showed some infiltration by mononuclear cells and greater variability in fiber size. The percentage of centronucleated fibers (CNF) in gastrocnemius, hamstring, and quadriceps muscles was counted in each of the three mouse models (Fig. 4b). The values were highest for the mdx4Cv/NRG and Sgca/NRG mice, but were also statistically significant for the BlAJ/NRG mice compared to NRG mice. In the BlAJ/NRG mice, a stronger effect was observed in hamstring muscle, compared to quadriceps and gastrocnemius, but all values were significantly elevated over the NRG control. However, CNF was significantly higher in mdx4Cv/NRG and Sgca/NRG mice relative to BlAJ/NRG mice.


Fig. 4: Hematoxylin and eosin staining and centronucleation.

(a) H&E staining in gastrocnemius muscle of NRG, BlAJ/NRG, Sgca/NRG, and mdx4Cv/NRG shows increases in the numbers of centronucleated fibers (CNF) in dystrophic mouse muscle. These data are quantified in (b). Scale bar = 75 um. Mice were six months old, and data are mean ± SEM with n = 4-9 with *p<0.05. G = gastrocnemius, Q = quadriceps and H = hamstring muscles.

Evans blue dye (EBD) staining was used to analyze membrane integrity. EBD is excluded from normal muscle fibers, but penetrates muscle fibers with perturbed membrane integrity, such as those from mdx4Cv mice 31, since the absence of dystrophin leads to disruption of the muscle membrane. After injection of EBD in our 6-month old mouse models, we evaluated the different amounts of EBD observed (Fig. 5a). All three of the mouse models demonstrated evidence of muscle pathology with EBD. In the gastrocnemius and quadriceps muscles, the mdx4Cv/NRG and Sgca/NRG mice both displayed significantly higher amounts of EBD relative to BlAJ/NRG and NRG controls (Fig. 5a). EBD in hamstring muscle was significantly higher in Scga/NRG than in the two other mouse models. The amount of EBD observed in BlAJ/NRG was significantly higher than in NRG mouse for hamstring muscle, but not for gastrocnemius and quadriceps muscle. We also noted an age-related increase in EBD in a study of BlAJ/NRG mice that ranged in age from 6-15 months; all three muscle groups showed significant elevations in EBD over age-matched NRG controls by 15 months of age (Fig. 6a).


Fig. 5: Evan’s blue dye, hydroxyproline, and treadmill studies.

(a) The three mouse strains are compared to NRG for penetration of Evan’s blue dye (EBD). mdx4Cv/NRG and Sgca/NRG mice showed elevated EBD values in all three muscle groups tested, whereas BlAJ/NRG mice exhibited greater EBD membrane permeabilization only in hamstring muscle compared to NRG mice. (b) Significant differences were observed in hydroxyproline deposits in gastrocnemius and hamstring muscles in mdx4Cv/NRG and Sgca/NRG mice, but not in quadriceps. BlAJ/NRG did not show significant increases at this age. (c) During treadmill exercise, BlAJ/NRG mice became exhausted significantly earlier than NRG mice, while mdx4Cv/NRG and Sgca/NRG mice became exhausted much earlier. Mice were six months old and data are mean ± SEM with n = 4-9, with *p<0.05. G = gastrocnemius, Q = quadriceps and H = hamstring muscles.

Furthermore, dystrophic mice have greater collagen deposition in muscle compared to wild-type mice 35. To characterize collagen deposition in our three mouse models, hydroxyproline of 6-month old mouse muscle was analyzed. We confirmed greater collagen deposition in the gastrocnemius and hamstring muscles of mdx4Cv/NRG mice compared to NRG mice (Fig. 5b). Collagen deposition in the Sgca/NRG mice was elevated to a similar degree (Fig. 5b). No significant difference of hydroxyproline was observed in quadriceps muscle between the three mouse models and the NRG control. The amount of hydroxyproline did not change between NRG and BlAJ/NRG mice in the six-month old mice (Fig. 5b). However, hydroxyproline was significantly elevated in the three muscles analyzed in older BlAJ/NRG mice (Fig. 6b), reflecting progressive pathology in the BlAJ/NRG animals.


Fig. 6: Characterization of aged BlAJ/NRG mouse model.

Whereas 6-month old BlAJ/NRG did not shown any statistical difference for EBD and hydroxyproline with NRG muscles, older BlAJ/NRG did. The amount of EBD (a) and hydroxyproline (b) were increased in the majority of the older BlAJ/NRG mouse muscles studied, compared to younger animals. Data are mean ± SEM with n = 3-6, with p<0.05. G = gastrocnemius, Q = quadriceps and H = hamstring muscles.

To evaluate muscle function, we used a treadmill assay. The mdx4Cv/NRG mice were able to run for dramatically less time than the wild-type NRG controls, confirming the functional deficit of the mdx4Cv/NRG mouse strain (Fig. 5c). For each of these assays, the well-established mdx/Scid strain was also evaluated, with the result that the new mdx4Cv/NRG mouse strain and mdx/Scid were similar in each case (data not shown). These results suggested that mdx4Cv/NRG mice were severely impaired in mobility and that placing the mdx4Cv mutation in the more severely immune-deficient background did not significantly alter disease pathology. The Sgca/NRG mouse showed similarly abbreviated treadmill times, compared to the NRG control, revealing a deficit in muscle function similar to that of mdx4Cv/NRG mice (Fig. 5c). We were also able to demonstrate a statistically significant deficit in the BlAJ/NRG mice with this assay (Fig.5c). This deficit was milder than that seen in mdx4Cv/NRG and Sgca/NRG mice, consistent with the milder phenotype of LGMD2B compared to DMD and severe forms of LGMD2D.


Immune-deficient mouse models are valuable resources for cell and gene therapy studies of genetic diseases. Genetic models allow assessment of phenotypes and detection of improvements to phenotypes that may be provided by experimental therapies. Such therapies often involve the use of foreign cells and genes whose use would be hampered in immune-competent animals. For example, the transplantation of human cells and the transfer of foreign genes stimulate rapid immune responses that would preclude longer-term experiments, since treated cells would be eliminated by the immune system. In recent years, the development of more completely immune-deficient mice that lack B, T, and NK cells that are less leaky than previous models such as Scid, have allowed long-term engraftment of human cells in strains such as NSG and NRG 28.

Crossing such mice with disease model mice remains a time-consuming and labor-intensive process. For that reason, we have made the mice described here available in a public repository. Although mdx4Cv/NSG and Rag2-Il2rb-Dmd mice were reported 32,33, they are not available in a public repository for easy access by the muscle community. Furthermore, the novel mdx4Cv/NRG mouse reported here may offer additional advantages for the study of muscle diseases. Because NRG mice are more resistant to irradiation, more flexibility in dose of irradiation can be used to eliminate satellite cells, without killing the mice. Less radioresistant mice can also be used for this purpose, using lower doses of irradiation to deplete host regenerative potential. This pre-conditioning creates space in the satellite cell niche for the enhanced engraftment of donor cells with regenerative capacity, assisting the evaluation of candidate therapeutic cells 29. The mdx4Cv/NRG mice reported here may thus be valuable for study of regenerative therapies for DMD. For LGMD2B, Scid/blAJ mice are available [Jackson Labs], but not more immune-deficient strains. For Sgca-/-, there is a lack of immune-deficient mice in public repositories. We have deposited the BlAJ/NRG strain described here to the Jackson Laboratories (Stock No: 029663, BlAJ/NRG, NOD.Cg-Rag1tm1MomDysfprmdIl2rγtmWjl/McalJ), as well as the mdx4Cv/NRG (Stock No: 030442, NOD.Cg-Rag1tm1Mom Dmdmdx-4Cv Il2rγtm1Wjl/McalJ) and Sgca/NRG (Stock No: 030443, NOD.Cg-Rag1tm1Mom Sgcatm1Kcam Il2rγtm1Wjl/McalJ) strains.

The severely immune-deficient mouse models for DMD, LGMD2B, and LGMD2D presented here will find use as key resources to test a variety of cell and gene therapies for these disorders. We validated that each of the mouse models carried the appropriate mutant alleles for the disease gene and immune deficiencies (Figs. 1-3). We also verified that each of the mouse strains was null for the disease gene of interest, using immunohistochemistry and western blots to show the absence of the relevant gene product (Figs. 1-3). To date, we have used the BlAJ/NRG mice in gene therapy experiments and have observed long-term retention (>3 months) of a luciferase transgene, which is otherwise rapidly rejected in immune-competent mice (J Ma, CP, HdB, MB, and MPC, manuscript submitted).

We performed characterization of the three mouse models to define their muscle phenotypes, using H&E staining and quantitation of the frequency of centronucleation as a measure of fiber turnover, Evan’s blue staining as a measure of sarcolemmal membrane integrity, and hydroxyproline quantitation as a measure of fibrosis. We also performed a treadmill assay on the mice to measure their ability to perform sustained running. These assays were carried out on mice of a similar age (~6 months), so that we could directly compare the level of pathology present in these three forms of muscular dystrophy disease model mice, although we note that because the lines are not completely inbred, there could be modifier genes that also impact the phenotypes observed. Consistent with the severe phenotypes associated with absence of dystrophin and sarcoglycans, significantly elevated values for the percent of centronucleated fibers were found in the mdx4Cv/NRG and Sgca/NRG mouse models (Fig. 4). These values likely reflect the degeneration and regeneration processes occurring in hind limb muscles as a result of the perturbation of the dystrophin-glycoprotein complex, in which both dystrophin and Sgca participate 1,2,3. Levels of centronucleation were also elevated in the BlAJ/NRG model, but to a lesser degree, consistent with the milder phenotype of dysferlin deficiency.

Evan’s blue dye penetration in six-month old mice was significantly elevated in all muscle groups tested in the mdx4Cv/NRG and Sgca/NRG mice, whereas the effect was predominant only in the hamstring muscle of BlAJ/NRG mice (Fig. 5a). Over time, in BlAJ/NRG mice 15 months old, Evan’s blue dye penetration became significantly elevated in all muscle groups tested (Fig. 6a). This finding is consistent with the progressive nature of muscular dystrophy and earlier involvement of upper limb muscles, which has been visualized previously in a LGMD2B model by luciferase live imaging 4. The collagen accumulation that accompanies the fibrosis process was detected in the gastrocnemius and hamstring muscles of mdx4Cv/NRG and Sgca/NRG animals by six months (Fig. 5b), but took longer to develop in BlAJ/NRG animals. These mice showed significant elevations of hydroxyproline in the upper limb muscles, quadriceps and hamstring, by 15 months of age (Fig. 6b).

NRG control mice were able to run for about 35 minutes under our assay conditions. We detected a significant deficiency in muscle endurance in the BlAJ/NRG mice, with average run time of about half this time. The mdx4Cv/NRG and Sgca/NRG mice were more severely affected, running for <5 minutes under these conditions. Again, the more severe phenotype was similar for both mdx4Cv/NRG and Sgca/NRG and significantly stronger than the effects in the BlAJ/NRG model (Fig. 5c). These characterizations provide useful baseline values that can be used to guide studies of therapeutic interventions when these mouse strains are utilized in preclinical studies of treatments that may be beneficial for these forms of muscular dystrophy.

Data Availability

Data are available from the Open Science Framework repository:

Competing Interests

The authors have declared that no competing interests exist.

Corresponding Author

M.P. Calos, Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305-5120, Tel. 650-723-5558; email

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The PJ Nicholoff Steroid Protocol for Duchenne and Becker Muscular Dystrophy and Adrenal Suppression Tue, 27 Jun 2017 11:00:26 +0000 Duchenne muscular dystrophy (DMD or Duchenne) is a progressive, life-limiting muscle-wasting disease that requires comprehensive, multidisciplinary care. This care, at minimum, should include neuromuscular, respiratory, cardiac, orthopedic, endocrine and rehabilitative interventions that address both the primary and secondary manifestations of the disease. The care needs of patients evolve over the cdourse of the disease and as they transition from childhood into young adulthood. In the past two decades, life expectancy has increased significantly by the use of corticosteroids and enhanced clinical management. Nevertheless, each year, patients with Duchenne muscular dystrophy are admitted to emergency departments and intensive care units where medical expertise thrives, but where expertise in rare diseases, such as Duchenne, may not. Emergency care for patients with Duchenne can be as complex as the disease process itself. While any illness or injury may occur in a person with Duchenne, some acute scenarios are much more common in the context of the disease. Making decisions about the clinical care of a person with Duchenne who presents with an acute illness can be quite difficult — in part, because of the extensive use of corticosteroids, which can lead to adrenal suppression. The life of a person with Duchenne needing emergency care may therefore depend upon the ability of the clinician on duty in the emergency department to recognize and mitigate adrenal suppression resulting from corticosteroid dependence. With this in mind, and drawing from expertise and experience with other steroid-dependent diseases, the ‘PJ Nicholoff Steroid Protocol’ was developed. The purpose of this protocol is to provide clinicians information regarding the safe management of corticosteroid during emergency situations in patients who may have accompanying adrenal suppression. The protocol explains how to recognize the signs and symptoms of acute adrenal crisis, how to prevent it with supplemental stress doses of corticosteroids, and how to taper doses after emergency care in order to prevent corticosteroid withdrawal.



Duchenne Muscular Dystrophy (DMD) is the most common and severe muscle disease presenting in childhood. It is caused by mutations in the dystrophin gene, located on the X chromosome, which causes a complete absence of dystrophin protein in muscle. Becker Muscular Dystrophy (BMD) is caused by partial absence of dystrophin; this disease is less severe and less common. As an X-linked disease, both diseases almost always affect males, though some females may be affected as well. Duchenne and Becker have a prevalence of 1 in 5000 males, in about a 2:1 ratio.1,2

People with Duchenne Muscular Dystrophy have progressive muscle weakness, which generally begins proximally and spreads to the legs, arms and other muscles. It is now known from a newborn screening study that the onset of disease is at birth, and probably in utero.3 Boys with Duchenne are born with serum creatine kinase (CK) values over 2000 U·L-1. The diagnosis is typically made between 3 and 5 years of age. Progressive muscle weakness leads to the loss of ambulation at 10 to 12 years old without treatment, though ambulation can be extended by one or two years through the use of corticosteroids. There is progressive respiratory muscle weakness, which leads to hypoventilation, respiratory insufficiency, and respiratory failure in adulthood. There is also a progressive cardiomyopathy leading to cardiac insufficiency. Prior to the use of corticosteroids, assisted ventilation and other interventions, the median life expectancy was age 19; however, today most deaths are due to heart disease and heart failure, and typically occur in the mid-20s, though survival into the 30s is no longer uncommon. 4,5

In the late 1980’s, it was established that corticosteroids could improve and prolong ambulation and slow the rate of decline in both Duchenne and Becker.6 The use of corticosteroids has also reduced the incidence and severity of scoliosis. However, chronic use of corticosteroids is associated with significant side effects that require monitoring and treatment.

International consensus recommendations on the comprehensive care for people living with Duchenne were published in Lancet Neurology in 2010. 5,7 A summary listing the key elements of Duchenne care was published in this journal in 2015.8

Given the multi-systemic nature of the disease, there should generally be an interdisciplinary team of care providers with whom to consult, including, at minimum, a coordinator, neurologist, doctor of physical medicine and rehabilitation, pulmonologist, cardiologist, social worker, dietician and primary care doctor. However, one issue in the management of Duchenne is that due to increased life expectancy, patients at some point need to transition out of pediatric care and on to adult care teams. Although the transition into adult care should be a process, clinicians in the emergency department should be aware that it is sometimes delayed or incomplete — and may in fact be triggered by the very acute illness or clinical event that brings them to the emergency room (ER).

While any illness or trauma may occur in a person with Duchenne or Becker, certain clinical scenarios are more common. These include exacerbations of the underlying respiratory or cardiac disease; kidney stones; and fractures as a result of low bone mineral density. Patients with Duchenne also seem to be prone to a particularly devastating consequence of fracture, fat embolism syndrome.9,10 Each medical crisis needs to be evaluated and responded to in the context of the expected complications of the disease or medications used for disease management — particularly the chronic use of corticosteroids, which can lead to suppression of the hypothalamus-pituitary-adrenal axis (HPA).

One danger is that, as pressing medical emergencies emerge, issues of daily medical care (including medications) may be pushed aside. However, the consequences of suddenly discontinuing or not appropriately dosing corticosteroids, particularly during a crisis can be quite severe, as the patient may have little to no ability to produce cortisol in response to stress. An additional issue for emergency departments is that many patients with Duchenne or Becker use Deflazacort, a corticosteroid that was recently approved in the United States but may be unfamiliar to doctors.

A recent case in point involved Phillip James “PJ” Nicholoff, a vibrant, 31-year-old man living with Duchenne muscular dystrophy. PJ had been treated with daily corticosteroids since the age of 6. He endured several pathologic fractures, likely a result of his steroid treatment and his non-ambulatory status. In November 2013, on a trip to Florida, he fractured his humerus and hip, and was transported by plane to a hospital closer to his home in the north. PJ had orthopedic surgery to manage both of these fractures. Following the surgery, however, he developed respiratory distress, tachycardia, hypotension, and later died. A review of the medical record suggested that he had not received appropriate stress-doses of steroids during his hospitalization. While PJ’s death may have been attributed to many causes, inadequate corticosteroid doses may have been a contributing factor.. Consequently, experts in the field held extensive discussions in order to develop a protocol on steroid dosing during an acute illness or other emergency.

The protocol addresses several issues:

  • How to define HPA suppression in a patient using corticosteroids
  • Appropriate corticosteroid stress doses for minor, moderate, and major stressors
  • Recommendations for corticosteroid withdrawal
  • How to test the HPA axis for continued suppression
  • Symptoms of acute adrenal crisis
  • Tests that can help diagnose adrenal crisis
  • Corticosteroid conversions/equivalent doses

The PJ Nicholoff Steroid Protocol

1. Background/Assessment

The normal basal secretion of cortisol from the adrenal gland is approximately 5-7 mg/m2/day in children or 8 -10 mg/day for adults.11,12 This amount increases during minor illnesses or surgery to approximately 50 mg/day (5x normal physiologic secretion), and typically return to baseline in 24 hours. Severe illness, trauma, or major surgical procedures cause increased cortisol production to about 75-150 mg/day (10x normal physiologic secretion), which return to baseline in about 5 days.13,14

Long-term administration of corticosteroids (primarily prednisone, prednisolone or deflazacort) may lead to suppression of the hypothalamic-pituitary-adrenal (HPA) axis, such that the adrenal glands no longer produce endogenous cortisol. Rapid reduction or abrupt withdrawal of corticosteroid therapy can cause secondary adrenal insufficiency and steroid withdrawal or deprivation syndrome, which may progress to adrenal crisis. Adrenal crisis is characterized by hypotension and hypovolemic shock, which may be life threatening.15 Recovery from suppression of the HPA axis after discontinuing corticosteroids can be prolonged (possibly 6 to 12 months) and may vary based on doses, dosing schedules and duration of corticosteroid therapy.14 Since there is a great deal of individual variability, it is not possible to predict with confidence which patients will be affected by adrenal suppression. Current practice is to administer supplemental (stress) doses of corticosteroids to patients with possible suppression of the HPA axis in the perioperative period and during acute illness to prevent acute adrenal insufficiency, or adrenal crisis.

2. Defining HPA Suppressed Patients:

Recommendations differ slightly in defining a suppressed patient, but general guidelines are as shown in table 1.16 These recommendations are primarily based on expert opinion and practice, as little data in this area have been published, but are based on the degree of medical/surgical stress and the likelihood of adrenal suppression.16 Consultation with an endocrinologist is recommended for questions or concerns.

Table 1: Defining Adrenal Suppression

Table 1: Defining Adrenal Suppression.16 Note: doses for prednisone and prednisolone are equivalent; conversion is required for patients taking deflzacort (5 mg prednisone or prednisolone = 6 mg deflazacort)

Patients receiving disease appropriate corticosteroid doses (at least 10 times above the physiologic cortisol dose) generally do not need stress doses if usual daily dose is continued. Patients who are on maintenance physiologic dose of hydrocortisone for primary disease of the HPA axis do require supplemental therapy.17 Recommendations for supplemental doses are generally divided by the severity of the stress the patient may experience (medical or surgical).

3. Corticosteroid Stress Doses for Patients Using Chronic Daily Corticosteroids

Table 2: Corticosteroid Stress Dosing

Table 2: Corticosteroid Stress Dosing

For patients using a high dose, twice-weekly corticosteroid*:

  • If patients are unable to take their usual corticosteroids by mouth due to nausea, vomiting, or npo status, they should take stress doses intravenously as indicated above.
  • During a moderate or major medical or surgical stressor, a cortisol level should be drawn prior to stress dosing.

4. Recommendation for withdrawal of Corticosteroid Therapy:

  • It is recommended that patients electing to discontinue the use of corticosteroids do so under the guidance of a neuromuscular provider and/or endocrinologist.

Weaning from corticosteroids, and reactivating the adrenal glands, may take several months to achieve. A recommendation for tapering chronic corticosteroids (generally managed in an outpatient setting) is as follows:

  • First, starting on a Monday, giving 20-25% lower corticosteroid dose for 2 weeks (or longer)
  • Next, if multiple daily doses are taken, start first to reduce to a single morning dose
    • Cut the dose 20-25% again for 2 weeks (or longer); continues this schedule
    • Continue until dose is near physiologic dose (3mg/m2/day of prednisone or 3.6mg/m2/day of Deflazacort)
  • When near physiologic dose, substitute corticosteroids with short acting form of corticosteroid or hydrocortisone (12 mg/m2/day of hydrocortisone)
  • This will also enable the patient to have a supply of hydrocortisone to be used for stress doses if needed in times of stress after coming off steroids
    • Continue to taper off by 20-25% each week (or longer)
    • Give every other day for 2 weeks (or longer)
    • Stop
    • Watch very carefully for signs of adrenal crisis (see below)
  • Alert patients and parents to signs/symptoms of adrenal crisis
  • If patients have symptoms of adrenal insufficiency during the taper, the patient should return to the previous steroid dose, which should be maintained longer

If the patient has a serious illness/injury during the taper, they may need a “stress dose” of corticosteroids:

  • Encourage parents to continue to report any serious events until 1 year post-taper
    • The stress doses of hydrocortisone is 30-50 mg/m2/day, or higher, for major stress (see Table 2)
    • Patients need to go to the emergency room if they are having signs or symptoms of adrenal crisis. Serum electrolytes with blood glucose and cortisol level should be obtained.
    • While it is appropriate to assess for acute adrenal crisis, this assessment should never delay treatment with a stress dose of hydrocortisone.
    • Patients should see an endocrinologist for evaluation of HPA axis during the process of corticosteroid therapy withdrawal.

Testing the HPA axis:18

  • After reaching half the physiological dose (5-6 mg/m2/day of hydrocortisone or 1-1.5 mg/m2/day of prednisone), monthly morning serum cortisol and ACTH should be assayed (may do less frequently), until they reach normal levels.
  • When baseline monthly morning serum ACTH and cortisol are normal, discontinue the corticosteroid and carry out the rapid ACTH stimulation test (may also be called cosyntropin, tetracosactide or Synacthen tests) monthly until post-stimulation cortisol response is normal (post-stimulus level > 20 mcg/dL). When this point is reached, it can be considered that the HPA axis has recovered.
  • It may be appropriate to recheck the rapid ACTH stimulation test after 1 year, to establish continued full recovery of the HPA axis.

Modification of above protocol:

  • Omit monthly AM cortisol and ACTH and perform an ACTH stimulation test in 3 months after discontinuation of corticosteroids
  • During this time (3 months before getting ACTH stimulation test), patients will need to take stress dose at the time of stress
  • If ACTH stimulation test result is abnormal (peak cortisol <20), patients will need to continue taking stress doses of hydrocortisone at the time of stress. (Patients should have a repeat ACTH stimulation test again in 1-2 months later and families would need to have teaching on this with an endocrine nurse.)

Alternatively, when laboratory tests cannot be carried out:

  • Patients who have used corticosteroids for prolonged periods can be considered as having suppression of the HPA axis up to 1 year after discontinuation of corticosteroid therapy and therefore need hydrocortisone stress dose coverage during the time of stress.

Risk factors for adrenal crisis include:19

  • Dehydration
  • Infection and other physical illness
  • Injury to the adrenal or pituitary gland
  • Missing usual doses of corticosteroids
  • Surgery
  • Trauma

Symptoms of adrenal crisis can include any of the following:15

  • Abdominal pain
  • Hypovolemic shock
  • Confusion or coma
  • Dehydration
  • Dizziness or light-headedness
  • Fatigue
  • Flank pain
  • Headache
  • High fever
  • Loss of appetite
  • Loss of consciousness
  • Severe hypotension
  • Nausea
  • Profound weakness
  • Tachycardia
  • Tachypnea
  • Slow, sluggish movement
  • Unusual and excessive sweating on face or palms
  • Vomiting

Exams and Tests

While it is appropriate to assess for acute adrenal crisis, this assessment should never delay treatment with a stress does of hydrocortisone.

Tests that may be ordered to help diagnose acute adrenal crisis include:

  • ACTH (cosyntropin) stimulation test
  • Cortisol level
  • Blood glucose
  • Serum potassium
  • Serum sodium
  • Serum pH

Table 3: Corticosteroid Conversion Table


This protocol, on how to manage patients with corticosteroid dependence, particularly during periods of stress and how to recognize and prevent an acute adrenal crisis, was named in honor of the late Philip James “PJ” Nicholoff, for his contribution to the global Duchenne community. Despite the tragic loss of PJ’s life, the “PJ Nicholoff Steroid Protocol” is a positive result that will impact the lives of people living with Duchenne and using corticosteroids around the world. We thank PJ and his family for encouraging the development of this resource.

A pdf of the PJ Nicholoff Protocol can be downloaded at: for the development of the PJ Nicholoff Steroid Protocol

In honor of the late Philip James “PJ” Nicholoff, for his contribution to the global Duchenne community.

  • Vincent’s Hospital, Indianapolis, IN
  • Philip Zeitler, Children’s Hospital Colorado, Aurora, CO
  • Sasigarn Bowden, Nationwide Children’s Hospital, Columbus, OH
  • Doug Biggar, Holland Bloorview Kids Rehab, Toronto, ON
  • Jerry R. Mendell, Nationwide Children’s Hospital, Columbus, OH
  • Anne M. Connolly, St. Louis Children’s Hospital, St. Louis, MO

Supporting Organizations

Parent Project Muscular Dystrophy (PPMD)

Parent Project Muscular Dystrophy (PPMD, is the largest nonprofit organization in the United States focused entirely on Duchenne. Started in 1994 by Pat Furlong, PPMD takes a comprehensive approach in the fight against Duchenne—funding research, raising awareness, promoting advocacy, connecting the community, and broadening treatment options. PPMD’s care objectives are to identify gaps in care for people with Duchenne and work toward solutions, and to work with clinicians and other health care professionals across the globe to ensure all Duchenne patients have access to optimal care.

Competing Interests Statement

The Authors have declared that no competing interests exist.

Corresponding Author

Kathi Kinnett (

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The C2A domain in dysferlin is important for association with MG53 (TRIM72) Mon, 05 Nov 2012 12:32:39 +0000


Dysferlin is a sarcolemmal protein, and dysferlin deficiency causes Miyoshi myopathy (MM) and limb girdle muscular dystrophy type 2B (LGMD2B) [1,2]. Based on the observation that dysferlin accumulates at wound sites in myofibers in a Ca2+-dependent manner, dysferlin is thought to mediate Ca2+-dependent sarcolemmal repair [3].

Mitsugumin 53 (MG53), also known as muscle-specific tripartite motif 72, is a recently identified protein involved in membrane repair in skeletal muscle [4]. Mice lacking MG53 suffer progressive myopathy [4], similar to dysferlin-null mice [3]. MG53 is localized in intracellular vesicles and plasma membranes in skeletal muscle, and it accumulates at injury sites in an oxidation-dependent, but not Ca2+-dependent, manner [4].

MG53 interacts with dysferlin and caveolin-3 to regulate sarcolemmal repair [5]. When expressed in C2C12 myoblasts that lack endogenous MG53, damaged membrane sites cannot be repaired in the presence of GFP-dysferlin, however, co-transfection of MG53 and GFP-dysferlin in these myoblasts results in GFP-dysferlin accumulation at injury sites [5]. These findings indicated that recruitment of dysferlin to the injury site of the membrane depends on MG53. However, it remains unclear whether the absence of dysferlin perturbs recruitment of MG53 to the injury site for membrane repair. A previous report has demonstrated the association of dysferlin with MG53 with co-immunoprecipitation (IP) assays using mouse skeletal muscle and C2C12 myoblasts transfected with dysferlin and MG53 [5]. However, which protein domains participate in this interaction between dysferlin and MG53 and whether this interaction is dependent on Ca2+ remain unclear. MG53 oligomerizes via disulfide bonds [4] and forms homodimers via a leucine-zipper motif in the coiled-coil domain [6]. The interaction between dysferlin proteins and MG53 monomers or oligomers has not been characterized in detail. To understand the precise role of dysferlin and MG53 in sarcolemmal repair, it would be helpful to determine whether dysferlin associates with MG53 monomers, oligomers, or both in a Ca2+-dependent manner.

Thus, to examine the biological role of the association between dysferlin and MG53, we used the following strategy to examine the effect of the absence of MG53 oligomers on dysferlin. We co-transfected mouse skeletal muscle with wild-type dysferlin-EGFP and RFP-tagged wild-type MG53 or a RFP-tagged MG53 mutant (RFP-C242A –MG53), and conducted a membrane-repair assay using a two-photon laser microscope. The C242A–MG53 mutant has been reported to form dimers, but not oligomers [6]. There is no report of simultaneous observation of dysferlin and MG53 during sarcolemmal repair; however, we have successfully performed real-time imaging of dysferlin-GFP and MG53-RFP after membrane injury in mouse skeletal muscle.

Dysferlin protein is absent or severely reduced in the skeletal muscle of patients with dysferlinopathy [7] and of SJL and A/J mice with mutations in the dysferlin genes [8]. To examine whether the absence of dysferlin affects the recruitment of MG53 to injury sites, we transfected skeletal muscle from dysferlin-deficient SJL and A/J mice with EGFP-MG53 and conducted membrane repair assays. These experiments are helpful in elucidating the molecular pathology of dysferlinopathy and revealed that MG53 accumulated in the skeletal muscles of dysferlin-deficient mice, which develop progressive muscular dystrophy.

We present evidence indicating that efficient sarcolemmal repair requires both dysferlin and MG53.


Immunoprecipitation. To examine the interaction between MG53 and dysferlin, mouse gastrocnemius muscles were lysed in lysis buffer containing 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1% NP-40, and Complete mini EDTA-free protease inhibitor cocktail (Roche) [9] supplemented with 1 mM CaCl2 or 2 mM EGTA. Lysates pre-cleared with Protein A/G agarose (Pierce) were incubated with polyclonal antibodies against mouse MG53 [4] or mouse dysferlin; the anti-dysferlin antibody was made in rabbit by injecting bacterial recombinant protein containing residues 1669 to 1790. The immunoprecipitated proteins were separated by SDS-PAGE and detected on immunoblots using the same antibodies used for IP or the anti-human dysferlin monoclonal antibody, NCL-Hamlet (Novocastra Laboratories).

A human MG53 cDNA was amplified by PCR and subcloned into pFLAG-CMV-4 (Sigma). Wild-type and truncated human dysferlin that were each tagged with c-myc were generated previously [10]. We also created five truncated human dysferlin constructs with the C2A domain (aa 1-149, 1-349, and 1-1080) and without the C2A domain (aa 130-2080 and 1081-2080). The sequence of each construct was verified by DNA sequencing. FuGENE 6 or E-xtremeGENE 9 (Roche) was used to transiently transfect COS-7 cells with MG53 and wild-type or mutant dysferlin constructs. Transfectants were cultured for 48 h and subsequently lysed in the same lysis buffer used to lyse mouse muscle, except that this buffer lacked CaCl2 and EGTA. Lysates pre-cleared with Protein G-Sepharose (GE Healthcare) were incubated with anti-FLAG (M2, Sigma) or anti-c-myc (9E10, Santa Cruz Biotechnology) monoclonal antibodies; Protein G-Sepharose was then added. Immunoprecipitated proteins were analyzed by immunoblotting using M2 and anti-c-myc polyclonal (A14, Santa Cruz Biotechnology) antibodies.

Pull-down assay. Fragments of the dysferlin C2A domain (corresponding to aa 1-129 of human dysferlin) were amplified as cDNA by PCR and subcloned into pGEX-5X-3 (GE Healthcare). Dysferlin p.W52R (TGG to CGG at c.527-529) and p.V67D (GTG to GAT at c.572-574) mutations were introduced by PCR using appropriate primers. GST fusion proteins expressed in BL21 E. coli were purified using sarkosyl [11] and bound to glutathione Sepharose 4B (GE Healthcare). COS-7 cells overexpressing FLAG-tagged human MG53 were lysed in lysis buffer containing 10 mM Na2HPO4, 1.8 mM KH2HPO4, 1% NP-40 (pH 7.4), 2 mM EGTA, various concentration of CaCl2, and Complete mini EDTA-free protease inhibitor cocktail. EGTA was used to chelate the free Ca2+ in solution and CaCl2 at various concentrations. The free calcium concentration was calculated using the free software CALCON3.6. Lysates were centrifuged to remove cellular debris, supplemented with 5 mM N-methylmaleimide (NEM) or 5 mM dithiothreitol (DTT), and finally subjected to protein cross-linking by treating with 2 mM glutaraldehyde (GA) for 5 min at room temperature, which was quenched with 100 mM Tris-HCl (pH 7.5) [6]. The cross-linked lysates were diluted with 75 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1% NP-40, 2 mM EGTA, various concentrations of CaCl2, and Complete mini EDTA-free protease inhibitor cocktail. Lysates pre-cleared with GST bound to glutathione Sepharose 4B were divided into aliquots and incubated with wild-type, p.W52R, and p.V67D dysferlin C2A-GST fusion protein bound to beads for 2 hr at 4°C. After three washes in lysis buffer containing 75 mM Tris-HCl (pH 7.5), 2× sample buffer (125 mM Tris-HCl (pH 6.8), 4% SDS, 20% (v/v) glycerol, and 0.004% bromophenol blue) was added to the beads, and the mixtures were incubated for 10 min at 85°C. Bound proteins were separated by SDS-PAGE and subjected to immunoblotting with the anti-FLAG antibody M2.

In vivo transfection and membrane repair assay. Twenty micrograms of N-terminal RFP-tagged human MG53 cDNA/pcDNA3.1 and/or C-terminal GFP-tagged human dysferlin cDNA/pcDNA3.1 plasmid DNA were injected into the flexor digitorum brevis of anesthetized, 4-week-old male C57BL6J and dysferlin-deficient SJL and A/J mice. Electroporation of plasmid DNA was performed using an electric pulse generator (CUY21SC, NEPAGENE) as described previously [12]. Seven days after electroporation, skeletal muscle myocytes (for whole-mount viewing) or individual myofibers were isolated and subjected to plasma membrane injury created by a two-photon laser microscope, LSM 710NLO with GaAsp Detectors (Zeiss) and Chameleon Vision II System (Coherent)[3]. Myofiber wounding using the 820-nm infrared laser and resealing analysis based on the kinetics and extent of FM1-43 or 4-46 dye (Molecular Probes) entry through open disruptions was carried out as previously described [3,13,14].

Ethics Statement. All experiments involving animals were performed according to the Procedure for Handling Experiments involving Animals of AIST (National Institute of Advanced Industrial Science and Technology) and approved by the Institutional Animal Care and Use Committee of AIST.


Association of MG53 and dysferlin in mouse skeletal muscle

We used an IP assay with protein from mouse muscle to confirm that endogenous MG53 associates with dysferlin in vivo. MG53 and dysferlin associated only in the absence of EGTA and CaCl2 (Fig. 1). The same result was obtained using C2C12 myotubes (data not shown). MG53 was specifically co-immunoprecipitated by the anti-dysferlin antibody, and conversely dysferlin was specifically co-immunoprecipitated by the anti-MG53 antibody. Thus, we confirmed that endogenous MG53 and endogenous dysferlin form a protein complex in mouse skeletal muscle without EGTA or CaCl2 supplementation.

Fig. 1: IP assay of dysferlin and MG53.

MG53interacts with dysferlin in mouse skeletal muscle. Extracts from wild-type mouse skeletal muscle were subjected to IP with polyclonal anti-MG53 antibodies or polyclonal anti-dysferlin antibodies. Immunoprecipitated proteins were subjected to SDS-PAGE and visualized on immunoblots treated with the same antibodies that were used for IP.

Identification of the MG53-associating domain of dysferlin

Next, we used IP to define the region of dysferlin that associates with MG53. Specifically, we used transient co-transfection to introduce a construct encoding full-length human MG53 tagged with FLAG and a construct encoding human dysferlin tagged with c-myc into COS-7 cells; for each co-transfection, full-length dysferlin or one of five deletion mutant forms of tagged dysferlin was used (Fig. 2). For deletion mutants that lacked the C-terminal domain of dysferlin, the transmembrane domain of dysferlin was retained to increase protein stability [10]. Transfectants were lysed in the same buffer that was used for IP assays of mouse skeletal muscle extract, except that this buffer lacked EGTA and CaCl2. Full-length dysferlin and deletion mutants that retained the N-terminal C2 (C2A) domain of dysferlin were co-immunoprecipitated by anti-MG53 antibody. In contrast, dysferlin mutants that lacked this N-terminal domain, Δ2-1080 and Δ2-129, failed to interact with MG53. These results indicated that the C2A domain of dysferlin was necessary for association with MG53.

Fig. 2: Identification of MG53-binding region of dysferlin.

The dysferlin C2A domain associates with MG53. Constructs encoding dysferlin deletion mutants were used for co-IP assays, and the results of these experiments are shown on the right. Deletion mutants encoding c-myc-tagged dysferlin mutants and FLAG-tagged full-length MG53 were co-expressed in COS-7 cells. IP and immunoblotting were performed using antibodies against the c-myc and FLAG tags. MG53 was co-immunoprecipitated with full-length dysferlin and the dysferlin mutants that lacked the C-terminus, but not with the dysferlin mutants that lacked the N-terminus.

Characterization of the association of dysferlin C2A domain with MG53 monomers and MG53 oligomer

C2 domains are known to bind to phospholipids and/or proteins in a Ca2+-dependent or Ca2+-independent manner [15]. Therefore, we used a pull-down assay to examine whether the Ca2+ concentration affected the association between MG53 and the dysferlin C2A domain. We used lysis buffer containing 75 mM Tris to reduce the change in pH that can result from the addition of CaCl2, to examine the calcium-dependency of the association between dysferlin and MG53. Reportedly, MG53 can exist as a monomer or an oligomer, depending on the redox state [4]. We used DTT for monomerization of MG53 by reducing sulfhydryl groups. Addition of 5 mM DTT resulted in complete dissociation of all MG53 oligomers (Fig. 3). To conduct a pull-down assay for MG53 oligomers, we treated cell lysates with an alkylating reagent, NEM, which reacts with sulfhydryl groups to form stable thioether bonds [6]. Multimers of MG53 were stabilized by chemical cross-linking with GA. Addition of 5 mM NEM to cell lysates resulted in oligomerization of MG53 (Fig. 3). In the presence or absence of Ca2+, MG53 oligomers associated with wild-type C2A-GST, whereas MG53 monomers did not associate with wild-type C2A-GST. In the absence of DTT or NEM, MG53 existed as oligomers including dimers, which associated with WT C2A-GST only in 10 mM free Ca2+ (Fig. 3, top).

Next, we generated two mutant versions of C2A-GST (W52R or V67D) to further characterize the association between MG53 and the C2A domain. A V67D missense mutation in the human dysferlin gene has been found in patients with MM and patients with LGMD2B [16]; similarly, the W52R dysferlin missense mutation has been found in patients with LGMD2B [17]. Each mutant C2A-GST, like the wild-type C2A, associated with MG53 oligomers when conditions included NEM in the presence or absence of Ca2+ (Fig. 3). However, the V67D mutation altered the calcium sensitivity of the association between C2A-GST and MG53 dimers; specifically, V67D-C2A-GST could associate with MG53 when conditions did not include NEM in the absence of Ca2+. In contrast, W52R-C2A-GST did not associate with MG53 when conditions did not include NEM in the presence or absence of Ca2+. These results revealed that the V67D mutation in the dysferlin C2A domain altered the Ca2+-dependence of the association between dysferlin and MG53 dimers.

Fig. 3: Pull-down assay of dysferlin C2A-GST and MG53.

COS-7 cells overexpressing FLAG-tagged MG53 were lysed and supplemented with DTT or NEM, and proteins in these lysates were cross-linked with GA. Cross-linked proteins were incubated with glutathione Sepharose 4B beads bound to wild-type C2A-GST, V67D C2A-GST, or GST. GST fusion proteins bound to beads were separated by SDS-PAGE, followed by Coomassie Brilliant Blue R-250-staining. Precipitated MG53 oligomers/monomers were detected on immunoblots using an anti-FLAG antibody. Mutations in the C2A domain affect the association of between dysferlin and MG53.

MG53 with a C242A missense mutation shows impaired accumulation at wound sites and attenuates the formation of dysferlin patches

To examine the biological role of the association between dysferlin and MG53 in sarcolemmal repair, we used mouse skeletal muscle co-transfected with dysferlin-EGFP and RFP-tagged wild-type MG53 or RFP-tagged mutant MG53 to perform a membrane repair assay. The mutant MG53 carried a C242A missense mutation and is designated RFP-C242A-MG53 here. MG53 with a C242A missense mutation reportedly exists as a monomer or dimer when expressed in mammalian cells, but does not form oligomers via disulfide bonding [4,6]. RFP-C242A-MG53 did not accumulate at wound sites as reported previously, and it was associated with defective sarcolemmal repair [4]. Co-expression of RFP-C242A-MG53 did not affect the subcellular localization of dysferlin in myofibers, and dysferlin was localized in a striated pattern (Fig. 4A). However, RFP-C242A-MG53 compromised the accumulation of dysferlin at injury sites (Fig. 4A, B). When the movement of dysferlin and wild-type MG53 were observed simultaneously in mouse skeletal muscle, RFP-wild-type MG53 accumulated more slowly at injury sites than dysferlin-EGFP (Fig. 4A). Accumulation of dysferlin-EGFP at wound sites stops within 5 seconds of injury and disperses gradually, while wild-type MG53 continues to accumulate for 200 seconds after injury (Fig. 4A and 4B).

Fig. 4: Membrane repair assay of myofiber transfected with dysferlin-GFP and RFP-MG53.

RFP-C242A MG53 perturbed the accumulation of dysferlin at wound sites in the sarcolemma. A. Dysferlin-GFP was simultaneously expressed with RFP-tagged wild-type MG53 or the RFP-C242A-MG53 mutant in mouse skeletal muscle. Arrowheads indicate sites of membrane injury, which were induced with a two-photon laser microscope. Dysferlin-GFP accumulated at the injury site in the presence of RFP-wild-type MG53, but no obvious accumulation of dysferlin-GFP was observed in the presence of the RFP-C242A-MG53 mutant. Scale bar, 10 mm. B.Time course fluorescence intensity (n=3) at wounded sites versus time. For every image taken, the fluorescence intensity of dysferlin-GFP at the site of the damage (circle of 5 mm in diameter) was measured with Zeiss LSM5 Image Examinar software. Data are means ± standard deviation.

MG53 accumulates normally at injury site of sarcolemma in dysferlin-deficient mice.

A previous study revealed that exogenous expression of MG53 in undifferentiated C2C12 cells was necessary for recruitment of GFP-dysferlin to sites of injury [5]. Conversely, to examine whether the recruitment of MG53 requires dysferlin, and to elucidate the molecular pathology of dysferlinopathy, we used skeletal muscle from dysferlin-deficient A/J mice transfected with EGFP-MG53 to perform a membrane repair assay. We confirmed that EGFP-MG53 accumulated at sites of injury (Fig. 5). Sarcolemmal repair was observed and confirmed by FM4-46-loading in A/J mice (data not shown). The accumulation of MG53 at the sarcolemmal wound was observed in A/L mice, similar to wild-type mice. Similar results were obtained from the membrane repair assay using dysferlin-deficient SJL mice.

Fig. 5: Membrane repair assay of myofiber using dysferlin-deficient myofiber transfected with GFP-MG53.

GFP-MG53 accumulated at sites of injury in the sarcolemma in dysferlin-deficient A/J mice, similar to wild-type mice. GFP-MG53 was expressed in wild-type or dysferlin-deficient A/J mice, and a membrane repair assay was performed using transfected myofibers. Subcellular localization of GFP-MG53 was similar between wild-type and A/J mice. Arrowheads indicate membrane injury sites, which were induced with a two-photon laser microscope. Scale bar, 10 μm.


Both dysferlin and MG53 are involved in membrane repair after injury in skeletal muscle. Dysferlin accumulates at wounded sarcolemmal sites, and this accumulation requires the influx of Ca2+ into the myofiber [3]. MG53 forms oligomers at the sarcolemmal injury site in an oxidation-dependent manner [4,6]. MG53 associates with dysferlin and facilitates vesicle trafficking to the site of membrane injury, and a recent finding suggests that MG53 and dysferlin may form a complex that participates in membrane repair in striated muscle [5]. To characterize the association between dysferlin and MG53, we used an IP assay and mouse muscle extract with or without exogenously added EGTA or CaCl2 to examine the Ca2+ dependency of this association. Using lysis buffer that lacked EGTA and CaCl2, we observed the association of dysferlin with MG53 in mouse skeletal muscle. Lysates lacking exogenously added EGTA and CaCl2 contain physiological concentrations of free calcium. Hence, low concentrations of calcium are likely to be necessary for the interaction between MG53 and dysferlin.

Our results indicated that MG53 oligomers associated with the dysferlin C2A domain in the presence or absence of Ca2+, whereas MG53 dimers associated with the dysferlin C2A domain in a Ca2+-dependent manner. We also revealed that pathogenic mutations in the dysferlin C2A domain (W52R and V67D) alter the association between this domain and MG53 dimers in a pull-down assay. In the absence of EGTA or Ca2+, dysferlin with a C2A missense mutation (W52R or V67D) did not associate with MG53 in an IP assay that used extracts from co-transfected COS-7 cells; however, full-length dysferlin with the most common pathogenic mutation found in Japan, a W999C missense mutation in the dysferlin domain, did associate with MG53 in these IP assays (data not shown). These results indicate that the dysferlin C2A domain is important for the association between dysferlin and MG53. Amino acid W52 in human dysferlin is located between the b5-sheet and the b6-sheet, and V67 is located in the b6-sheet [18]. Both residues are reportedly important for the C2 structure, particularly those of the b-sheet, and are predicted to coordinate calcium [18].

Recently, MG53 was reported to form homodimers, which are essential for MG53-mediated sarcolemmal repair [6]. We used pull-down assays to investigate associations between MG53 monomers or MG53 dimers and the dysferlin C2A domain, and we found that MG53 dimers associated with dysferlin in a Ca2+-dependent manner. An increase in the cytoplasmic Ca2+ level is necessary for dysferlin accumulation at wounded sarcolemmal sites [3]. The intracellular Ca2+ level is maintained at 50-100 nM in resting mammalian cells, but this increases to 6 μM after membrane puncture in Swiss-3T3 cells [19]. The influx of extracellular Ca2+ through the wound site is required for vesicle fusion with the plasma membrane and formation of a repair patch in skeletal muscle, but MG53 trafficking to the wound site does not require Ca2+ [4]. In pull-down assays in the present study, we demonstrated a selective association between the wild-type dysferlin C2A domain and MG53 dimers at a free Ca2+ concentration of 10 μM, but not at lower or higher free Ca2+ concentrations. These findings indicated that the concentration of free Ca2+ is important for association of dysferlin with MG53 dimers, and suggest that MG53 dimers not only form oligomers, but also associate with dysferlin in response to sarcolemmal injury. The altered Ca2+ sensitivity of the association between dysferlin with a mutation in the C2A domain and MG53 dimers in the pull-down assay also suggested that the C2A domain was important in the Ca2+-dependent association between dysferlin and MG53 dimers.

We were able to analyze the movement of dysferlin and MG53 in real time during sarcolemmal repair in a membrane repair assay that employs mouse myofibers that express dysferlin-EGFP and RFP-MG53. This is the first report to demonstrate that dysferlin and MG53 have different accumulation patterns at wound sites, and this result indicated that dysferlin and MG53 have different functions in sarcolemmal repair. Our studies also revealed that MG53 carrying a C242A missense mutation can suppress the accumulation of dysferlin at the wound site; this finding, together with results from pull-down assays, suggests that MG53 dimers play an important role in sarcolemmal repair.

Our studies also revealed that MG53 accumulated at injury sites in the sarcolemma in dysferlin-deficient mice, similar to wild-type mice. However, dysferlin-deficient SJL and A/J mice have a progressive muscular dystrophy phenotype, suggesting that MG53 is necessary but not sufficient for efficient sarcolemmal repair.

Competing Interests

The authors have declared that no competing interests exist.


Address for correspondence : (C. Matsuda)

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The proteasomal inhibitor MG132 prevents muscular dystrophy in zebrafish Mon, 21 Nov 2011 09:29:49 +0000


The zebrafish Danio rerio , has rapidly been adopted as an organism of choice for all aspects of the drug discovery pipeline [ [1] , [2] , [3] ]. We have therefore developed a zebrafish medium-throughput platform as a test bed for therapeutic advancement for muscular dystrophies. The zebrafish system offers unique advantage for drug screening in a vertebrate model organism, and in particular muscular dystrophies are especially amenable due to their early, robust and readily recognisable phenotypes [ [4] , [5] ]. Their small size, embryonic status, low cost and ease of drug delivery directly via the water make zebrafish a very attractive model for whole organism screening. Zebrafish show a typical vertebrate development pattern, and in the mutants, perturbation of muscle architecture and muscle function is readily observable even in the embryonic stages [ [4] , [5] , [6] ]. In addition, of the genes known to be mutated in human forms of muscular dystrophy, all but one are represented in the zebrafish genome and those investigated so far exhibit dystrophic phenotypes in zebrafish [ [7] , [8] ]. Although candidate compounds identified in fish would need to be validated in mammals before being taken on to human therapy, the low cost and speed of candidate drug screening, far outweighs any disadvantages.

A recent screen from the Kunkel Group has also validated this approach and identified a number of compounds that appear effective in reducing dystrophic symptoms in zebrafish [9] , in particular PDE5 inhibitors appear to be useful in this regard as they have also been shown to be effective in mdx mice [ [10] , [11] ]. Previous studies from the Lisanti group and ourselves suggested that tyrosine phosphorylation of dystroglycan is an important mechanism for controlling the association of dystroglycan with its cellular binding partners dystrophin and utrophin, and also as a signal for degradation of dystroglycan [ [12] , [13] , [14] ]. The Lisanti group further demonstrated that inhibition of the proteasome was able to restore other dystrophin glycoprotein complex (DGC) components in both mdx mice that lack dystrophin and in explants of DMD patients [ [15] , [16] ]. We have therefore chosen to examine the proteasomal inhibitor MG132 as a proof of principle in the zebrafish system comparing wildtype with dystrophic sapje larvae, which have a premature stop codon in the dystrophin gene, express no full-length dystrophin protein and exhibit a dystrophic phenotype [6] .



Heterozygous sapjet222a zebrafish [6] were maintained using standard procedures. Pairs of heterozygous sapje fish were mated, and embryos collected and raised at 28.5°C. Embryos 24 hours post fertilisation (hpf) were transferred in groups of 20 into 12-well plates containing 1ml of E3 media. Embryos were exposed to the proteasomal inhibitor MG132 (Calbiochem) at various concentrations from a DMSO stock diluted in E3 media, for 48 hours at 28.5°C. The final concentration of DMSO was 1% in both treated and control wells.


At 3dpf embryos were anaesthetised in tricaine, and viewed between polarising filters on a dissecting microscope. The proportion of fish with an abnormal muscle birefringence, as determined by eye, was counted. For quantitative analysis, images were captured using a SPOT camera. Quantification of muscle damage was measured by taking a line scan from the 5th somite after the head along 1mm of the dorsal region of the somites in the direction of the tail. Line scans, L(i), in which i represents gray-scale intensities over the length of the sample, were subjected to standard Fourier analysis [17] and the resulting transforms, given as power spectra, were tested for significance by one-way ANOVA (Bonferroni-test) at selected spatial frequencies. In the Fourier analysis, the line scans (>2,000 points) were converted to mm-scale, divided into 50% overlapping stretches and windowed with a Blackman-Harris 4-term window [18] each giving seven to nine 250-points long samples. Thus, we obtained 7-9 spectral samples, which were averaged to improve the estimates of their power spectra, <|L(f)|2>, where | | denotes the norm, f spatial frequency and <> the average over the different stretches.

Ethics Statement

No specific ethics approval under UK and EU guidelines was required for this study, as all zebrafish used were less than 5.2dpf, and are therefore not protected under the Animals (Scientific Procedures) Act. Embryos were obtained from adult zebrafish by a regulated procedure under the UK Home Office project licence number 40/3134. Adult zebrafish are maintained in UK Home Office approved facilities in the Medical Research Council Centre for Developmental and Biomedical Genetics aquaria at the University of Sheffield.

Results and Discussion

Using birefringence as a marker of muscle damage, compared to wildtype larvae sapje zebrafish exhibit a typical mottled appearance (Figure 1AB), indicative of disruption to the muscle architecture and a dystrophic phenotype. The birefringence phenotype is not readily visible until around 72hpf, but in order to test compounds that may prevent the onset of muscular dystrophy, drugs must be added before fish can be phenotyped. Sapje is a recessive allele and behaves in a normal Mendelian manner. Consequently experimental procedures are carried out on a mixed population of fish comprising 25% wildtype, 50% heterozygote and 25% homozygote sapje . As can be seen from Figure 1B, the extreme dystrophic phenotype of the homozygote sapje fish is clearly visible. However potential treatments that may be beneficial, might not completely restore the normal muscle architecture and give a regular birefringence pattern as seen in Figure 1A. We therefore examined potential methods to quantify the extent of muscle damage (or recovery) in dystrophic sapje embryos. Simple quantification of the brightness of the birefringence was found to be unreliable for two reasons. Birefringence is very orientation dependent and if embryos are not aligned in precisely the same way, relative to each other, position dependent changes in birefringence result, which have nothing to do with changes in muscle structure. Furthermore, simple brightness over the whole fish cannot distinguish between a fish with some dark and some bright somites, such as in Figure 1B, or a fish with a low level of muscle damage giving a more even but generally reduced birefringence (data not shown). We therefore chose to use line scanning and Fourier analysis to more precisely quantify the muscle damage. Figures 1CD, represent line scans of the individual fish shown in Figures 1AB, with the yellow bar representing the position of the scan. Wildtype fish have a regular and even pattern of birefringence, with brighter peaks in the somite and darker troughs corresponding to the myotendinous junctions. Sapje fish however, show a very irregular pattern of peaks and troughs due to the disruption of the birefringence pattern induced by the lack of dystrophin (Figure 1D). Fourier analysis of multiple wildtype or sapje fish is shown in Figure 2. Compared to wildtype, which at 3dpf have a single well defined intensity frequency of 10/mm corresponding to the observable somite boundaries, sapje fish have a completely disordered frequency distribution, with major peaks at 4 and 10/mm but many other intervening frequencies representing the very broken birefringence pattern in these fish. Statistical analysis of these frequency plots reveals a clear statistical difference (p=0.029) comparing as few as 4 wildtype with 6 mutant fish. This analysis provides a straightforward quantification method to determine the extent of muscle damage in zebrafish.

Fig. 1: Birefringence images of wildtype (A) and sapje (B) zebrafish at 3dpf.

Yellow bar shows the position of the line scan used to generate the intensity profiles shown in C and D for wildtype and sapje fish respectively.

Fig. 2: Line scans and corresponding Fourier analysis from wildtype (A,B) and 3dpf sapje (C,D).

Left-hand plots (A,C) show the raw individual line scans for each fish used. Right-hand graphs (B,D) show the Fourier transforms of the data. Wildtype fish have a single frequency of 10/mm representing the actual number of somites per mm in these fish. sapje fish, whilst showing an underlying frequency of 10/mm, due to the disruption to the birefringence caused by the lack of dystrophin, also exhibit several other smaller frequency modes. As is clear to the eye, pixel power of wildtype and mutant fish populations are significantly different. At 4 cycles/mm, WT: 748 ± 536 and mutant: 36504 ± 26159 (mean ± SD, p = 0.029; one-tail ANOVA).

To validate further the sapje model we used the proteasomal inhibitor MG132, which has previously been shown to have beneficial effects in mdx mice [ [19] , [20] ]. Treatment of sapje zebrafish with MG132 significantly reduces the number of dystrophic fish with an aberrant birefringence pattern from ~25% to ~15%, a rescue of approximately 40%. Titration of MG132 levels indicated a concentration-dependent restoration of birefringence in sapje zebrafish compared to DMSO vehicle alone. Expressed as percentage rescue of the 25% dystrophic population, the effect of MG132 reached a plateau at around 2µM with an EC50 of 0.4µM (Figure 3). This effective concentration range is an order of magnitude lower than doses reported to be effective in the hindlimb of mdxmice [20] or in explants from BMD and DMD patients [19]. This may however be attributable to the relative permeability of zebrafish embryos to water borne agents and therefore reflects the ease of delivery and effective dose achieved in the tissue, rather than any real difference in efficacy in fish as compared to mammals.

Fig. 3: Dose response curve for the effect of MG132 in rescuing the dystrophic phenotype in sapje zebrafish.

Each data point represents the percentage rescue of the dystrophic phenotype, taking the proportion of dystrophic fish in vehicle alone (1%DMSO: 0 µM MG132) as 0% rescue. Data are mean ± SEM of three independent experiments, following administration of MG132 or vehicle from 1dpf to 3dpf. EC50 = 0.4µM, maximal effective dose 2µM.


This study adds further to the utility of zebrafish as a model of choice for testing muscular dystrophy therapeutics. In addition to the previous study validating the efficacy of PDE inhibitors [9] we can now add proteasomal inhibitors. Furthermore, zebrafish have also been demonstrated to be suitable for the testing of exon skipping strategies to treat muscular dystrophy [21] , making them an invaluable part of the toolkit for the evaluation of a range of muscular dystrophy therapies.

Competing Interests

The authors have declared that no competing interests exist.

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