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. 2008 Nov 11;17(1):19–25. doi: 10.1038/mt.2008.239

Gene-mediated Restoration of Normal Myofiber Elasticity in Dystrophic Muscles

Stefania Puttini 1, Małgorzata Lekka 2, Olivier M Dorchies 3, Damien Saugy 1, Tania Incitti 4, Urs T Ruegg 3, Irene Bozzoni 4, Andrzej J Kulik 5, Nicolas Mermod 1
PMCID: PMC2834969  PMID: 19002166

Abstract

Dystrophin mediates a physical link between the cytoskeleton of muscle fibers and the extracellular matrix, and its absence leads to muscle degeneration and dystrophy. In this article, we show that the lack of dystrophin affects the elasticity of individual fibers within muscle tissue explants, as probed using atomic force microscopy (AFM), providing a sensitive and quantitative description of the properties of normal and dystrophic myofibers. The rescue of dystrophin expression by exon skipping or by the ectopic expression of the utrophin analogue normalized the elasticity of dystrophic muscles, and these effects were commensurate to the functional recovery of whole muscle strength. However, a more homogeneous and widespread restoration of normal elasticity was obtained by the exon-skipping approach when comparing individual myofibers. AFM may thus provide a quantification of the functional benefit of gene therapies from live tissues coupled to single-cell resolution.

Introduction

Muscular dystrophies are a group of inherited diseases characterized by the wasting of muscle tissue and progressive muscle weakness. They are most often caused by mutations in genes involved in muscle structure and function.1,2,3 In particular, Duchenne muscular dystrophy is caused by mutations in the dystrophin gene. Although several therapeutic approaches are being developed to restore the expression of proteins such as dystrophin, quantitative assessment of the functional recovery of muscle fibers remains difficult.

Dystrophin, a large protein expressed primarily in skeletal muscles, can be divided into three distinct regions: (i) the N-terminus having actin-binding domains, (ii) 24 spectrin-like triple-helical coiled-coil repeats, and (iii) the C-terminal region containing motifs involved in protein–protein interactions.4 Dystrophin is part of a flexible link between the actin cytoskeleton and the extracellular matrix through a membrane-associated protein complex and it thus contributes to the mechanical stability of the muscles. Hence, a major role for dystrophin is the protection of the sarcolemma from the shearing forces and mechanical stress associated with muscle contraction. The absence of sufficient functional dystrophin in muscle fibers results in disruptions of the cell membrane, necrosis and loss of functions.5,6

Several approaches have been developed to restore the expression of dystrophin or of its functional homologue utrophin.7 Utrophin is expressed in many tissues and cell types in the body. In skeletal muscle, utrophin is abundantly expressed during the embryonic life and it is progressively replaced by dystrophin thereafter. Utrophin is overexpressed in the skeletal muscles of Duchenne muscular dystrophy patients as in the mouse, dog, and cat animal models, presumably as part of a compensatory mechanism.8,9 Dystrophin and utrophin both bind the cytoskeleton through a similar actin-binding domain and they associate with the components of the dystrophin–glycoprotein complex that spans the membrane and connects to extracellular proteins.7 Several studies have shown that utrophin can replace dystrophin functionally, and that elevated utrophin expression improves muscle strength in dystrophic animals.10 Because of its widespread expression, utrophin has been proposed as an alternative therapeutic strategy for Duchenne muscular dystrophy, as dystrophin may elicit an immune response in dystrophic patients.

The expression of full-length utrophin or dystrophin remains difficult with viral vectors, because of their very large coding sequence. However, expression of shorter derivatives such as the mini- or micro-dystrophins yielded significant muscle function restoration.7 Alternatively, small nuclear RNAs that mediate exon skipping were used to remove a nonsense mutation-containing exon from the dystrophin mRNA. These chimeric antisense small nuclear RNAs proved to be very effective in restoring dystrophin synthesis both in human Duchenne muscular dystrophy cells,11 and in the mouse mdx model through local or systemic injection of adeno-associated viral (AAV) vectors.12,13 Moreover, body-wide and stable expression of the antisense molecule, as well as dystrophin expression rescue and functional benefits, were shown to persist for almost the entire life span of the animal.14

A main symptom of muscular dystrophies is the weakening of muscles, as exemplified by a reduced resistance to lengthening contractions. This may confer a distinct resistance pattern in response to an external deforming force such as that applied by atomic force microscopy (AFM).15 AFM has been used to measure the stiffness of single cells in culture and of in vitro differentiated myotubes,16,17,18 or to distinguish normal myocardial tissue from its infarcted fibrotic counterpart.19 However, its potential ability to evaluate the therapeutic effect of gene transfer approaches using live tissues has not been evaluated. In this study, we show that the elastic properties of individual muscle fibers can be probed by AFM in conditions that preserve the anchorage of live myofibers to their extracellular matrix environment, using muscle explants from healthy and dystrophic mice. This approach provided an easy and sensitive detection of the mechanical properties of dystrophic and normal myofibers and of the changes elicited by the expression of utrophin or dystrophin. AFM may thus provide a promising approach to probe some of the physiological properties of live tissues in response to molecular or cellular therapies.

Results

Although progress has been made toward gene or cell therapies of muscular dystrophies, assays allowing the demonstration of a functional recovery of the treated muscle fiber beyond the expression of the transgene remain difficult to achieve. The establishment of functional assays that may be applied to small samples such as murine muscles or biopsies of human muscles would thus be advantageous. In this study, we first evaluated whether AFM may be adapted to probe the elastic properties of live myofibers within explants of murine muscles.

Comparison of the elastic properties of normal and dystrophic muscles

We first addressed the ability of AFM to detect physiologically relevant differences from normal and dystrophic muscle fibers in conditions resembling as much as possible their natural tissular environment. Therefore, we assayed thick slices of muscles explants rather than isolated fibers or primary cells in culture. The tibialis anterior (TA) muscles of normal and dystrophic mice were collected and cut longitudinally into 1-mm-thick sections to preserve the integrity of the myofibers. These explants were then immobilized on microscope glass slides and submersed in cell culture medium (Figure 1a), and 100–700 individual measurements were performed on the muscle sections to probe the local properties of representative numbers of single muscle fibers. The mechanical properties of the muscular tissues were then determined on the basis of force versus displacement measurements using an indentation depth of 500 nm (Figure 1b), following a method described previously.17,20

Figure 1.

Figure 1

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Principles of AFM measurements of muscle resistance to deformation. (a) Scheme of the experimental AFM setup. The tendons of muscle sections glued to a glass coverslip, immersed in DMEM medium, and placed in the “liquid cell” setup of the AFM scanner. (b) Principles of the quantification of muscle elastic properties. Only curves recorded during the approach of the AFM probe to the muscle surface are shown. The subtraction of two curves obtained from one recorded on the glass substrate (showing negligible deformation) from that recorded on the muscle explant, results in the force-versus-indentation dependence used for the determination of the Young's modulus value describing in a quantitative manner the muscle's ability to deform elastically. AFM, atomic force microscopy.

To determine whether AFM is sufficiently sensitive to detect elasticity changes elicited by the lack of dystrophin, we compared several independent explants of TA muscles from normal and mdx dystrophic mice. Plotting revealed a significantly lower resistance of mdx muscles to reversible deformation (Figure 2a,b, filled bars). Fitting of the expected normal distribution with Gaussian curves allowed the calculation of the Young's modulus average value, which is a measure of the stiffness of the biological sample in response to an applied load. When compared directly, the muscles of normal mice yielded Young's modulus values of ~4.2 ± 1.1 kPa on average, whereas those of the dystrophic mice were about threefold lower (1.4 ± 0.4 kPa). The lower modulus values obtained from the dystrophic muscle explants indicate lower resistance to deformation, as may be expected from the lack of dystrophin.

Figure 2.

Figure 2

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Effect of cytochalasin D on muscle stiffness. Normalized Young's modulus value distributions performed on thick longitudinal sections of (a) wild-type (NTOT = 126) or (b) dystrophic (NTOT = 150) mice muscle (black bars) are shown in comparison with those obtained from an adjacent section of the same muscle treated with cytochalasin D (gray bars, NTOT = 192 and NTOT = 168, respectively). The distributions were fitted with Gaussian curves, where the centers of distribution denote the Young's modulus mean value.

Effect of cytochalasin D on the mechanical properties of normal and dystrophic muscles

The mechanical properties of individual myofibers may be affected by their interaction with the surrounding extracellular matrix and with the other fibers, as well as by the resistance of their cytoskeleton. To assess whether the Young's modulus values directly reflect the action of dystrophin as a physical bridge between the cytoskeleton and the extracellular matrix, muscle explants were treated with cytochalasin D. This compound binds to the positive extremity of the cytoskeletal F-actin filaments, which inhibits polymerization and leads to the disruption of the cytoskeleton.21 Cytochalasin D is thus commonly used to measure the contribution of actin filaments to the cellular stiffness.16 Assay of adjacent sections of muscles treated with cytochalasin D decreased the Young's modulus of both normal and mdx muscle fibers to similar values (0.8 ± 0.3 kPa versus 1.3 ± 0.7 kPa; Figure 2, gray columns). This indicates that disruption of the cytoskeleton abolishes most of the difference in the stiffness of wild-type and dystrophic muscle explants.

Overall, these findings support the notion that AFM may be adapted to probe the elasticity of the sarcolemmal and sub- sarcolemmal membrane compartment within live myofibers, which may provide a quantitative assessment of the physical continuity of the membrane with the cytoskeleton, and, therefore, of proper dystrophin function.

Mechanical resistance of utrophin-expressing mdx muscles

To assess whether overexpression of utrophin may compensate for the lack of dystrophin and restore normal levels of resistance to pressure, a utrophin expression vector was electroporated in vivo in the TA muscle of normal and mdx mice using a constant-current device. The distribution of transgene expression was assessed by green fluorescent protein (GFP) fluorescence, as expressed from a co-electrotransferred vector. Fluorescence pictures showed nonhomogenous distributions of GFP expression, with ~30–80% of the fibers displaying high levels of GFP fluorescence, depending on the area investigated (Figure 3a,b). The electrotransfer of the utrophin expression vector resulted in an approximately twofold increase of the overall utrophin content of dystrophic TA muscles, and moderate extra-synaptic utrophin labeling was observed throughout the plasmalemma, as expected from utrophin overexpression (see Supplementary Figure S1a,b). Consecutive muscle sections enriched in GFP-expressing fibers were prepared and assayed using AFM. Sections from the corresponding contralateral muscle injected with the GFP vector only were used as controls.

Figure 3.

Figure 3

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Localization of muscle fibers expressing utrophin and effects on muscle function. Fluorescence microscopy analysis of mouse TA muscle after electrotransfer of utrophin and GFP expression vector plasmids was performed on (a) a whole muscle or on (b) a transversal muscle section. Muscle regions showing the brightest area were sectioned longitudinally and assayed by AFM as described in the Materials and Methods section. The Young's modulus value distribution of control male mdx muscles are shown in c (NTOT = 252, black columns), whereas gray columns illustrate the distribution of utrophin-expressing muscle sections (NTOT = 186). The latter distribution shows two contributions: unaffected values corresponding to those of dystrophic muscle fibers (31% of recordings) and values elevated after electrotransfer (69%). Phasic responses were recorded under isometric conditions from electrically stimulated TA muscle of dystrophic mice after in vivo electroporation of plasmids encoding GFP alone (MDX-GFP) or GFP and utrophin (MDX-Utro), as displayed in d. TA muscles from normal mice after electroporation of the GFP plasmid alone (C57-GFP) were used for comparison. Traces represent average from six to nine muscles per group after normalization to the muscle cross-sectional surface. For clarity, only one error bar every 10 ms is shown. Note that the specific force of GFP-expressing dystrophic TA is significantly lower than that of its normal counterpart, and that the introduction of the utrophin plasmid into the dystrophic TA muscle fully restored force. AFM, atomic force microscopy.

The histogram obtained for contralateral control samples of 9-week-old male mdx mice showed one maximum as before, yielding a Young's modulus value of 0.8 ± 0.4 kPa when fitted with a Gaussian function (black bars and solid line in Figure 3c). Similar experiments performed on age-matched female mdx mice yielded a similar distribution of the Young's modulus values (1.0 ± 0.4 kPa; see Supplementary Figure S2a). Upon utrophin overexpression, the distribution became noticeably wider, showing a second broad peak (Figure 3c, gray bars). The first maximum likely results from the populations of muscle fibers not expressing utrophin, because the average Young's modulus is close to the value obtained for nontreated mdx muscles. The second Gaussian curve revealed fibers having Young's modulus values close to those of normal fibers. This second population likely results from utrophin-mediated recovery of a normal muscular stiffness in the subset of myofibers which incorporated the utrophin expression vector upon in situ electrotransfer.

The distribution was fitted with two Gaussians following the procedure described in Materials and Methods (Figure 3c, dashed lines), and the area under each fitted curve was used to estimate the percentage of muscle fibers belonging to a given population. Approximately 31% of the assays showed abnormally low Young's modulus values indicating insufficient or no utrophin expression. The remaining 69% of muscles fibers showed stiffness values around 4 kPa, close to the 4.2 kPa observed from normal muscles (compare Figures 2a and 3c), indicating that utrophin expression may have led to a full functional restoration of the sarcolemmal resistance of dystrophic fibers.

Recovery of some of the muscular functions upon utrophin expression was assessed 4 weeks after in vivo electroporation under deep sedation.3,22 Electrically elicited isometric tensions were recorded from whole TA muscles in situ, in conditions preserving innervation and blood supply. Isometric force measurements were performed with male dystrophic mice electrotransfered with a mixture of both GFP- and utrophin-expression vectors, and on the contralateral muscle electroporated with the GFP expression vector alone. Normal C57BL/6J mice electrotransfered with the GFP vector in both legs were used as reference. The phasic twitch responses were corrected for the mean cross-sectional surface of the muscles (Figure 3d). The specific force developed by GFP-expressing dystrophic muscles was significantly lower than the one of their GFP-expressing normal counterparts, in agreement with previous measurements of noninjected muscles of normal and dystrophic mice.3 The introduction of utrophin-expressing vector into dystrophic TA muscle corrected the specific phasic force output to values similar to those of normal muscles. In addition, utrophin expression in dystrophic muscle resulted in the correction of the time required to reach 80% relaxation form tetanus, as assessed after repetitive tetanizations of the TA muscles. Of note, the rates for contraction and relaxation were significantly increased in utrophin-electroporated muscles (Figure 3d), indicating that utrophin may induce significant changes in the function of dystrophic muscles and/or that it may favor the occurrence of slow fiber types. In addition, utrophin expression did not yield full recovery of all muscle contractile characteristics, as it normalized phasic but not tetanic twitch tension, and it did not elicit a normal resistance to repetitive tetanization (Supplementary Figure S3 and data not shown).

Persistence and specificity of the increased resistance of utrophin-expressing mdx muscles

From a therapeutical point of view, a gene-mediated therapy should provide the patients with long-lasting effects to limit repetitive transgene administrations. Therefore, we investigated the time course of the effects of utrophin expression upon electrotransfer in 5-week-old male mdx mice (Figure 4a–f). One week after electrotransfer the amount of utrophin-expressing fibers was 81% (Figure 4b), and this value decreased moderately to 63 and 69% 1 and 2 months after electrotransfer, respectively (Figure 4d,f).

Figure 4.

Figure 4

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Time course of utrophin-mediated mdx muscle improved stiffness. The TA muscles of male mdx mice were explanted and assayed by AFM 1 week after the electrotransfer of either (a) the GFP expression vector alone (NTOT = 480), owr of both the GFP and (b) the utrophin-expression vectors (NTOT = 705). Muscles treated as for the a and b were assayed 1 month (c, NTOT = 443 and d, NTOT = 680) and 2 months (e, NTOT = 402, and f, NTOT = 690) after vector electrotransfer. AFM, atomic force microscopy.

Potential effects of utrophin expression were also assayed on normal muscles. Although changes of the elastic properties could also be detected from normal muscle fibers overexpressing utrophin, they were much smaller than those noted when assaying mdx muscles both in the number of altered fibers as well as in the range of variation of Young's modulus values (Supplementary Figure S4a,c).

Interestingly, smaller changes of muscle fiber elasticity were also elicited by the expression of GFP alone in mdx mice. One week after electrotransfer, this effect was visible in 36% of the mdx fibers (Figure 4a) and it was still observed in 11 and 17% of the fibers, 1 month and 2 months, respectively, after electrotransfer of the GFP vector (Figure 4c,f). This nonspecific effect was not seen after GFP expression in normal muscles (Supplementary Figure S4a,c), nor upon an electrotransfer process performed without the GFP expression vector (Figure 5a). This indicates that the nonspecific increase of muscle stiffness resulted from the expression of GFP per se in mdx muscles, but not from the in vivo electroporation procedure. The molecular nature of this effect is not known, but it may be due to GFP interacting with the actin-binding domain of myosin.23 In contrast, expression of utrophin without GFP yielded a normalization of the elastic properties of mdx muscles as noted before, indicating that it results from a specific effect of utrophin (Figures 4 and 5c).

Figure 5.

Figure 5

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Effect of utrophin and dystrophin expression on muscle function as probed by AFM. The TA muscles of male mdx mice were explanted and assayed by AFM either (a, c) 1 month after electrotransfer of the utrophin-expression plasmid alone (c, NTOT = 364) or after mock electrotransfer performed in the absence of expression vector (a, NTOT = 504), or 3 months after transduction with an small nuclear RNA (snRNA)-AAV viral vector–mediating dystrophin RNA exon skipping (d, NTOT = 374) or with the nontransduced contralateral controls (b, NTOT = 290). AFM, atomic force microscopy.

AFM assay of the rescue of dystrophin synthesis by exon skipping

Next, we investigated a distinct gene therapy approach based on the removal of a stop codon in exon 23 of the dystrophin mRNA by an exon-skipping mechanism.14 A chimeric small nuclear RNAs carrying sequences complementary to the dystrophin exon to be skipped was expressed from an AAV vector-injected intramuscularly in the TA muscle. Expression of dystrophin in treated mdx muscles was confirmed by western blot analysis, whereas it was undetectable in nontreated dystrophic muscles (Supplementary Figure S1c). AAV vector-injected or nontreated muscles were collected 3 months after injection of the viral vector, and muscle sections were generated and processed for AFM analysis. The myofibers Young's modulus values were increased in the transduced muscle as compared to control, and their distribution was homogeneous and unimodal (Figure 5b,d). The Young's modulus value of the treated dystrophic muscle fibers was increased to 3.9 ± 1.4 kPa, reaching values not significantly different from those of wild-type muscle fibers (4.2 ± 1.1 kPa). In addition, the variability of the Young's modulus values was not significantly more heterogeneous than those of the normal muscle (compare Figure 2a with Figure 5c,d). Finally, fibers showing no improvement, i.e., possessing Young's modulus values ~1 kPa, were virtually absent from dystrophin-AAV treated muscles. These findings correlate well with previous observations showing that >80% of TA muscle fibers express dystrophin in treated mdx mice.14 On the other hand, this contrasts with results obtained from electrotransfer of the utrophin vector, which yielded abnormally elevated or absent effects in some measurements, following the heterogeneous transgene expression ranging from undetectable expression to very high levels (compare Figures 3b and 5c). Finally, the AAV-mediated exon-skipping treatment allowed the recovery of normal and homogeneous resistance values in nearly all TA muscle fibers. Thus, AFM may be used to quantify expression homogeneity of myofibers.

Discussion

The muscle weakening associated with dystrophies may be expected to modify the elasticity of the muscle fibers before any loss of function occurs. Consistently, myotubes differentiated in vitro from dystrophic progenitor cells were previously shown to display lower stiffness values when compared to those of normal animals.24 However, opposite effects were reported elsewhere,18 and a reduced stiffness was also attributed to a partial loss of actin filaments and/or microtubules.25 Thus, whether the lack of dystrophin per se may affect myofiber stiffness remained unclear, and the assay of the stiffness of dystrophic muscle fibers treated to express dystrophin or utrophin had not been reported. In this study, we show that the elastic properties of dystrophic and normal tissues can be easily distinguished, and that normal functional properties can be restored by the expression of dystrophin or utrophin in dystrophic muscles.

Conditions were chosen to investigate the properties of individual fibers within the context of muscle tissue sections. An indentation depth of 500 nm was chosen to assess alterations in the organization of actin filaments just beneath the cell membrane and their connections with the dystophin–glycoprotein complex that spans the sarcolemma. The observed changes in Young's modulus values were comparable to those noted previously on myotubes differentiated in vitro using myoblasts obtained from dystrophic or normal mice,24 and the ratio of fibers displaying normal Young's modulus values paralleled closely the extent of gene transfer, thus comforting our conclusion that single myofibers are discerned within the tissue explants.

AFM provided a very sensitive detection of the dystrophic phenotype before the appearance of overt symptoms, and when little defects can be detected in normal muscle contraction conditions. In addition, it allowed the differentiation of the efficacy of distinct therapeutic strategies. In vivo electroporation of the full-length utrophin-coding sequence yielded variable gene transfer efficiencies and elasticity values when comparing distinct myofibers, and utrophin overexpression is a likely cause of the occurrence of unusually high Young's modulus values for some of the fibers. In contrast, dystrophin expression rescue by the viral vector and AFM results were more homogeneous. Indeed, even if the small nuclear RNA had been overexpressed, it would not be expected to lead to abnormally elevated dystrophin levels, owing to its exon-skipping effect on the endogenous RNA.

Myofiber resistance to deformation was corrected to a similar extent with either approach, as expected from the functional replacement of dystrophin by utrophin, and the normalization of muscle elasticity correlated well with the improvement of isometric contractile properties. Thus, expression of utrophin in a subset of muscle fibers sufficed to induce an essentially complete restoration of muscle tension, as found with the dystrophin-expressing mdx muscles. However, slower kinetics of contraction and relaxation were noted, suggesting a phenotypic adaptation of the muscle to utrophin expression, while dystrophin expression rescue was distinctly associated with increased contraction kinetics.14 This may reflect a different biological function of the embryonic protein as compared to dystrophin.

Thick muscle sections as used here allowed the assay of live myofibers within an environment that resembles that of an intact tissue. Up to 1,000 individual AFM measurements can be performed daily and this assay may complement current studies of single-muscle fibers, where the interactions mediating tissue cohesion are lost, or whole muscle strength recordings, where the contribution of individual fibers cannot be distinguished. This study indicates that AFM may allow the presymptomatic diagnosis of dystrophies of unknown genetic basis or the evaluation of the efficacy of molecular therapies from small biopsies of muscles which strength cannot be assessed readily, and it paves the way to the evaluation of therapies for other diseases that affect tissue morphogenesis or cell structure.

Materials and Methods

Muscle electrotransfer with GFP and utrophin-expressing vectors. All procedures involving mice were performed in accordance with institutional guidelines and authorizations for animal research. In vivo experiments were carried out on 6–8-week-old C57BL6 mice (Iffa Credo, Charles River, France) or age-matched mdx5Cv mice bred in-house.26 The TA muscles were injected with 0.8 U/µl of bovine hyaluronidase in 25 µl of 0.9% NaCl solution under isofluorane anesthesia.27 Two hours later, plasmid DNA (1 µg/µl, 30 µl) in 0.9% NaCl solution was injected intramuscularly. All injections were performed under Ketaminol (100 mg/kg) and Narcoxyl (10 mg/kg) sedation. Control muscles were injected with 30 µg CMV-GFP plasmid only and test muscles were injected with 15 µg of CMV-GFP and/or 15 µg CMV-utrophin (kindly provided by A. Briguet and Santhera Pharmaceuticals). Following the intramuscular injection of DNA, a three-needle-electrode was positioned over the lower hind limb, and 52 ms pulses of 0.1A constant current were applied using an electroporator for in vivo use, according to the manufacturer's instructions (VGX Pharmaceuticals, The Woodlands, TX). Alternatively, the skin was removed and spatula-shaped electrodes were applied on each side of the TA muscle and eight 200 V pulses were applied as described previously.28 Samples were collected 7, 28, or 56 days after the injection of plasmids. The efficiency of electrotransfer (i.e. in vivo electroporation) was monitored from GFP fluorescence as expressed from a co-injected vector. Immediately after muscle excision, whole muscles were visualized on a fluorescence microscope with a green fluorescence filter at room temperature. Muscle sections from normal or mdx mice were collected from the regions where GFP fluorescence was the brightest for AFM assays when GFP was expressed, while slices were randomly picked in other cases.

Utrophin expression and localization assays. After isometric force measurements, the TA muscles were cut at mid-length. The distal parts were embedded in tragacanth gum and frozen in liquid nitrogen-cooled isopentane. The proximal parts were snap-frozen in liquid nitrogen. Samples were stored at −80 °C until processed for determination of utrophin expression by immunostaining and western blotting, respectively. Immunolabeling was performed on 10 µm-thick transverse sections using the M.O.M kit (Vector laboratories, Burlingame, CA) according to the manufacturer instructions on the use of mouse antibodies on mouse tissue. Nonspecific binding sites were blocked for 1 hour, then the sections were incubated 2–3 hours with a mouse antiutrophin antibody diluted 1/1,000 (clone DRP2, Novocastra laboratories, Newcastle Upon Tyne, UK), extensively rinsed, incubated 1 hour with an AlexaFluor594-conjugated goat anti-mouse antibody, incubated 5 minutes with Hoechst dye no. 33258 (bis-benzimide, 1 µg/ml), extensively rinsed again, and finally mounted with Mowiol (Calbiochem, San Diego, CA). Pictures were captured with a SpotInsight digital camera (Visitron Systems, Puccheim, Germany) installed on a Zeiss 200M microscope equipped for epifluorescence.

Dystrophin-AAV vector administration to mdx mice. Clone U1#23 and AAV vector preparation were as described previously.13 Two 6-week-old male mdx mice were administered with the dystrophin-AAV9 vector as described previously (4.3 × 1012 genome copies/ml) via direct intramuscular injections.14,28 The injection volumes were 45 µl (split over three sites) in the right TA muscle. Animals were killed 3 months after injection, and injected or uninjected muscles were explanted for AFM measurements. For western blot analysis, proteins were extracted from powdered tissue with 100 mmol/l Tris HCl pH 7.4, 1 mmol/l EDTA, 2% SDS, and a protease inhibitor cocktail (Roche Diagnostics, Basel, Switzerland). Fifty micrograms of proteins were loaded onto 6% polyacrylamide gels, electrophoresed and transferred to Nitrocellulose transfer membranes (Protran, Schleicher & Schuell, Sanford, ME). Western analysis was performed as described.13

Sample preparation for AFM. Immediately after the killing, the TA muscles from normal or dystrophic mice were collected and immersed in DMEM medium (Dulbecco's modified Eagle's medium; Invitrogen, Carlsbad, CA). Afterward, each muscle was cut longitudinally (to preserve the integrity of the myofibers) into 1-mm thick sections and dimensions ~6 × 2 mm essentially as described.18 Sections were kept in DMEM and measurements were performed the same day. When noted in the text, muscle slices of normal or mdx mice were treated with 0.1125 µmol/l of cytochalasin D in DMEM for 30 minutes at 37 °C in a cell culture incubator.16 Afterward, the samples were washed with DMEM alone and immediately assayed with AFM. A muscle slice was removed from the medium and glued onto a glass coverslip using two 0.5 µl droplets of cyanoacrylate adhesive (AXIA, Kitta, Korea) placed at both extremities of the muscle slice (Figure 1a). The slice was immediately immersed in DMEM and AFM measurements were performed. Muscle explants were not kept in vitro for >10 hours.

AFM and Young's modulus determination. AFM assays were performed using a commercially available device (PSIA XE120, Park Systems, Korea) equipped with the “liquid cell” setup. Standard silicon nitride, nonsharpened, gold coated cantilevers (MLCT-AUHW, Veeco), with the nominal spring constant value of 0.01 N/m were used in all measurements. The deflection of the cantilever was measured using the laser–photodiode technique. The coverslip holding the muscle slice immersed in DMEM was placed into the “liquid cell” placed on the AFM scanner. For a given condition, around NTOT = 100–700 different locations were measured. The force acting between the end of the tip and the investigated surface causes cantilever deflection. By monitoring this deflection as a function of the relative sample position, one can record the force applied versus the relative distance between the probe tip and the investigated surface, the so-called force curve (Figure 1b). By comparing the experimental force curve (measured on a muscle slice, dashed line) with the corresponding calibration curve (taken from the reference glass sample, solid line), the relation between the load force applied to the sample and the resulting indentation depth can be obtained. Such dependence i.e., force‐versus‐indentation curve, is characteristic for each material and can be used for the quantification of elastic properties of the investigated sample by calculating the Young's modulus. Its value is determined on the basis of Sneddon's description of the elastic half-space behavior pushed by a hard axisymmetrical indenter.29 Thus, the applied load force F depends on the indentation depth Δz according to the formula:

graphic file with name mt2008239e1.jpg

where Em is the Young's modulus of a single-muscle fiber, ηm is the Poisson coefficient assumed to be 0.5 as muscle fibers can be treated as noncompressible material, and α is the opening angle of a probing cone (the AFM four-sided pyramidal tip was approximated by a cone with α = 35°).19 As the thickness of muscles section lying on the glass coverslip was much larger than the 500 nm indentation depth taken for calculation, the influence of the hard glass surface was negligible.

To check the applicability of the Hooke's law in the whole range up to Fmax, the difference between the load F500 and the maximum load force Fmax was calculated and plotted as a function of the maximum load Fmax applied in the AFM measurements (Supplementary Figure S5). The points placed above the line indicate the data influenced by the presence of hard substrate while those below the line results from the inappropriate determination of the contact point, i.e., the point where the cantilever touches the surface. Therefore, only the points located around the line were chosen for further analysis. To avoid potential disruption of the myofibers membrane by the AFM cantilevel tip, soft cantilevers with a large radius of curvature (~50 nm) were used with a maximum load force of about 1 nN. Absence of puncture of myofiber membrane by the AFM tip was confirmed experimentally as described previously.2,30

To determine the mean value of the Young's modulus, the modulus distribution was fitted with the Gaussian function. The center of the distribution corresponds to the mean value while half of its width, taken at half height, denotes the standard deviation. In histograms obtained from muscles expressing utrophin, two contributions were visible: the first one revealing Young's modulus similar to those of nonelectroporated mdx muscles and the other one showing an effect of the utrophin. The number of fibers expressing utrophin was estimated as the difference between (i) the whole histogram (approximated by a sum of one or two Gaussian curves) and (ii) the population of unaffected muscle fibers (fitted with a single Gaussian corresponding to the first maximum).

Isometric force recordings. Isometric force recordings were performed essentially as described previously,13,21 with minor adaptations for small-sized muscles. Animals were anesthetized by i.p. injection of a mixture of urethane (1.5 g/kg) and diazepam (5 mg/kg). The TA muscle was exposed by opening the overlying skin. The adherences of the half-distal part to the surrounding tissues were gently cut away to minimize artifacts during force development. Care was taken not to disturb innervation and blood supply. The distal tendon was released and linked to a force transducer coupled to a custom-made LabView interface for trace acquisition and analysis. The knee joint was firmly immobilized. Two thin electrodes made of 0.15 mm diameter copper wire were inserted into the TA muscle, perpendicular to the long axis and equidistant from both ends of the muscle. The muscle was electrically stimulated with 0.5 ms pulses of controlled intensity and frequency via a custom-build stimulator. After setting the optimal muscle length and optimal current intensity, three to five phasic twitches were recorded to measure the absolute phasic twitch force (Pt), the time to peak, and the time for half relaxation from the peak (RT1/2). After a 3-minute pause, the muscle was subjected to a force–frequency test, where the force was recorded using 200 ms bursts of increasing frequency (from 10 to 100 Hz by increments of 10 Hz) with one burst every 30 seconds. Complete tetanus was usually achieved at 80–90 Hz. The strongest response was taken as the absolute optimal tetanic force (Po). Finally, after another 3-minute pause, the resistance to repetitive tetanization was measured: 60 stimuli were delivered, each consisting of a 1-second titanic contraction and a 3-second rest. The maximal tension was usually obtained during the first 5 stimulations. The amplitude of the response decreased as the stimuli were repeated. The time for 80% relaxation from tetanus (RTT80) was determined from the first 30 tetani. The same procedure was repeated on the second leg. The leg to be measured first was randomly assigned at the time of anesthesia. The absolute phasic and tetanic forces (in mN) were converted into specific force (mN/mm2 of muscle section) after normalization for the average TA cross-sectional area. The cross-sectional area (in mm2) was determined by dividing the TA muscle mass (in mg), by the product of optimal muscle length (in mm) and by the density of mammalian skeletal muscle (d = 1.06 mg/mm3).

Supplementary MaterialFigure S1. Determination of utrophin and dystrophin expression levels.Figure S2. Young's modulus values of TA muscles from female mdx mice.Figure S3. Effect of utrophin expression on whole muscle strength.Figure S4. Effects of GFP and utrophin expression on the Young's modulus values of normal muscles.Figure S5. Linear regime of muscle deformation in the AFM assay.

Supplementary Material

Figure S1.

Determination of utrophin and dystrophin expression levels.

Click here for supplemental data (367.5KB, tiff)
Figure S2.

Young's modulus values of TA muscles from female mdx mice.

Click here for supplemental data (215.1KB, tiff)
Figure S3.

Effect of utrophin expression on whole muscle strength.

Click here for supplemental data (894.1KB, tiff)
Figure S4.

Effects of GFP and utrophin expression on the Young's modulus values of normal muscles.

Click here for supplemental data (474.8KB, tiff)
Figure S5.

Linear regime of muscle deformation in the AFM assay.

Click here for supplemental data (239.4KB, tiff)

Acknowledgments

We are grateful to ParkSystems for the free loan of an XE 120 AFM system, to A. Khan, R. Draghia-Akli, and VGX Pharmaceuticals for help with in vivo electroporation, to A. Briguet and T. Meyer for the gift of plasmids, and to O. Patthey-Vuadens for expert immunofluorescence studies. This work was supported by grants from the Swiss Foundation for Research on Muscle Diseases to N.M. and U.T.R., the European CLINIGENE Network of Excellence (N.M.), the Swiss National Science Foundation (U.T.R.), the Telethon and Duchenne Parent Project Italia (I.B.), the Universities of Lausanne and Geneva, and the Institute Pasteur Cenci-Bolognetti.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1.

Determination of utrophin and dystrophin expression levels.

Click here for supplemental data (367.5KB, tiff)
Figure S2.

Young's modulus values of TA muscles from female mdx mice.

Click here for supplemental data (215.1KB, tiff)
Figure S3.

Effect of utrophin expression on whole muscle strength.

Click here for supplemental data (894.1KB, tiff)
Figure S4.

Effects of GFP and utrophin expression on the Young's modulus values of normal muscles.

Click here for supplemental data (474.8KB, tiff)
Figure S5.

Linear regime of muscle deformation in the AFM assay.

Click here for supplemental data (239.4KB, tiff)

Articles from Molecular Therapy: the Journal of the American Society of Gene Therapy are provided here courtesy of The American Society of Gene & Cell Therapy

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