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Vol. 73, No. 8 August 2014 pp. 789Y797
ORIGINAL ARTICLE
Glycosaminoglycan Modifications in Duchenne Muscular Dystrophy: Specific Remodeling of Chondroitin Sulfate/ Dermatan Sulfate Elisa Negroni, PhD, Emilie Henault, MSc, Fabien Chevalier, PhD, Marie Gilbert-Sirieix, PhD, Toin H. Van Kuppevelt, PhD, Dulce Papy-Garcia, PhD, Georges Uzan, PhD, and Patricia Albanese, PhD
Abstract Widespread skeletal muscle degeneration and impaired regeneration lead to progressive muscle weakness and premature death in patients with Duchenne muscular dystrophy (DMD). Dystrophic muscles are progressively replaced by nonfunctional tissue because of exhaustion of muscle precursor cells and excessive accumulation of extracellular matrix (ECM). Sulfated glycosaminoglycans (GAGs) are components of the ECM and are increasingly implicated in the regulation of biologic processes, but their possible role in the progression of DMD pathology is not understood. In the present study, we performed immunohistochemical and biochemical analyses of endogenous GAGs in skeletal muscle biopsies of 10 DMD patients and 11 healthy individuals (controls). Immunostaining targeted to specific GAG species showed greater deposition of chondroitin sulfate (CS)/dermatan (DS) sulfate in DMD patient biopsies versus control biopsies. The selective accumulation of CS/DS in DMD biopsies was confirmed by biochemical quantification assay. In addition, highperformance liquid chromatography analysis demonstrated a modification of the sulfation pattern of CS/DS disaccharide units in DMD muscles. In conclusion, our data open up a new path of investigation and suggest that GAGs could represent a new and original therapeutic target for improving the success of gene or cell therapy for the treatment of muscular dystrophies.
From the INSERM U972, Paul Brousse Hospital, Villejuif (EN, FC, GU); and EAC CNRS 7149, CRRET Laboratory, Sciences and Technology Faculty, Paris-Est Creteil University, Cre´teil (EH, FC, MG-S, DP-G, PA), France; and Department of Biochemistry (280), Nijmegen Center for Molecular Life Sciences, Radboud University Nijmegen Medical Center, Nijmegen, the Netherlands (THVK). Send correspondence and reprint requests to: Elisa Negroni, PhD, INSERM U972, Paul Brousse Hospital, 12 Avenue Paul Vaillant Couturier, Villejuif 94807, France; E-mail: [email protected] Present address: Elisa Negroni, UMRS 974, INSERM, Universite´ Paris 6, CNRS UMR 7215, Institut de Myologie, Baˆtiment Babinski, Groupe Hospitalier Pitie´-Salpeˆtrie`re 47, boulevard de l’Hoˆpital, 75651 Paris Cedex 13, France. Georges Uzan and Patricia Albanese contributed equally to this work. This study was supported by grants from Association Fran0aise contre les Myopathies and Region Ile-de-France. Elisa Negroni was supported by a fellowship from Association Fran0aise contre les Myopathies. Fabien Chevalier was supported by a doctoral fellowship from Region Ile-de-France. All authors declare no conflicts of interest.
Key Words: Chondroitin sulfate, Dermatan sulfate, Duchenne muscular dystrophy, Extracellular matrix, Fibrosis, Glycosaminoglycans.
INTRODUCTION Duchenne muscular dystrophy (DMD) is the most common inherited muscular dystrophy of childhood, affecting 1 in 5,000 live male births (1). It is caused by mutations in the dystrophin gene, leading to the absence of a functional dystrophin protein (2). In DMD patients, widespread skeletal muscle degeneration most often leads to progressive muscle wasting, cardiac dysfunction, respiratory failure, and, ultimately, death in the third decade of life (3). Dystrophin links the extracellular matrix (ECM) to the cytoskeleton of muscle fibers through the dystrophin-associated glycoprotein complex (4); it is a cytoskeletal protein essential for the structural integrity of the membrane of myofibers (i.e. the sarcolemma) (5). Absence of dystrophin results in permanent fragility and leakiness of the sarcolemma, which lead to increased calcium influx and progressive degeneration and necrosis of muscle fibers with subsequent loss of muscle tissue and progressive fibrosis (6). Fibrosis appears early in muscles of DMD patients, increases with age, and represents a major hurdle hampering the success of gene or cell therapies at advanced stages of the disease (7, 8). Indeed, fibrosis is a complex process characterized by excessive accumulation of ECM components in which there is progressive replacement of functional muscle tissue by nonfunctional connective tissue, leading to worsening of the disease and irreversible loss of muscle function. In normal tissue, the ECM is a dynamic structure that has important roles in cell signaling and cell anchorage, thus maintaining tissue homeostasis (9). It is mainly composed of structural glycoproteins and fibrous proteins such as collagens and proteoglycans (PGs) (10). Proteoglycans are the most abundant components of the nonfibrillar ECM. They are composed of a protein core to which long, linear, highly sulfated glycosaminoglycan (GAG) chains are covalently at tached. Proteoglycans display different functions that are principally mediated by the GAG chains (11). Glycosaminoglycans are polymers of alternating disaccharide residues. Their structure is highly complex owing to the nature of repeating disaccharide units, the heterogeneity of glycosidic linkage, sulfation patterns, and hydrophobic regions, which
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regulate their interactions with surrounding molecules (12, 13). Studying GAG signaling is complex because their biologic functions are directly related to their fine structure. Moreover, the structural variability of GAG sequences is generated during their biosynthesis under the control of multiple enzymes that progressively assemble and simultaneously modify the original polysaccharide backbone (12). The 2 main sulfated GAG families in muscle are heparan sulfate (HS) and chondroitin sulfate (CS)/dermatan sulfate (DS) (14, 15). They are involved in a variety of physiologic processes through activation, binding, or protection of various heparin-binding proteins (HBPs), such as growth factors, chemokines, cell surface receptors, and other ECM components that regulate key steps in development and tissue homeostasis (14Y16). Many studies have focused on the core proteins of PGs in skeletal muscle, irrespective of the GAG chains they harbor. For example, the role of syndecan (HS/CS-PG) core proteins has been studied in muscle development and regeneration, where they regulate satellite cell activation and myofiber formation (17). The roles of decorin (CS-PG) and biglycan (CS-PG) core proteins have been investigated in mdx mice in which these PGs have been shown to be upregulated (18, 19). In human DMD muscle, these CS-PGs appear to be altered, depending on the cohort of patients analyzed (20, 21). None of the studies describing changes in PG spatiotemporal expression during myogenesis or muscular dystrophy have focused on GAG structural epitope remodeling, which is directly related to the interaction with HBPs. In the present study, we performed immunochemical and biochemical analyses of endogenous GAGs in DMD patient and control human muscle biopsies to investigate changes in spatial distribution and structure that may contribute to the pathophysiology observed in this devastating muscle disease.
MATERIALS AND METHODS Patients Human skeletal muscle biopsies from 10 DMD patients (aged 8Y16 years) and 11 healthy individuals (controls [CTR]) were analyzed. Characteristics of the biopsies are summarized in the Table. After informed consent had been provided by the participants, all biopsies were obtained from Myobank (a partner in the European Union network EuroBioBank) in accordance with French and European Union legislation on ethical rules.
Muscle Histology Muscle biopsies were mounted on gum tragacanth (6% in water; Sigma-Aldrich, St Louis, MO) and snap-frozen in liquid N2-cooled isopentane. Staining was carried out on 5-Km-thick transverse cryosections. For the assessment of tissue morphology and visualization of fibrosis and connective tissue, muscle sections were stained with hematoxylin and eosin and Sirius red and examined by light microscopy. Fibrosis was quantified and expressed as a percentage of the surface of the whole crosssectional area on Sirius redYstained slides using Image J 1.44o analysis software (http://imagej.nih.gov/ij).
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Immunofluorescence Immunostaining was performed on 5-Km-thick cryostat muscle sections, as previously described (22). The following primary antibodies were used: antiYneonatal myosin heavy chain (NNmyoHC) (1:50, rabbit polyclonal) (23) and antispectrin (1:50, NCL-Spec1, mouse monoclonal IgG2b; Novocastra). Sections were then incubated with appropriate secondary antibodies, and nuclei were stained with Hoechst (Sigma-Aldrich). To visualize GAGs, we performed immunostaining, as previously described (24). Briefly, cryosections were fixed with 4% paraformaldehyde, incubated for 10 minutes with 50 mmol/L NH4Cl, and saturated with 3% bovine serum albumin/ phosphate-buffered saline for 45 minutes. Glycosaminoglycans were then stained overnight at 4-C with different phage display antibodies: AO4B08 (1:20), which recognizes HS species and was described to interact with an ubiquitous N-, 2-OY, and 6OYsulfated saccharide motif (25); IO3H10 antibody (1:5), which is described to recognize chondroitin 6-O-sulfate (26) and was used to reveal CS; and LKN1 antibody (1:5), which is described to recognize GlcA2S-GalNAc4S motif (27) and was used to stain DS. Immunostainings were carried out overnight at 4-C. Bound antibodies were detected with mouse antiYvesicular stomatitis virus monoclonal antibody (P5D4, 1:200) then incubated with a secondary antibody coupled to Alexa Fluor 555 GAM (1:200; Life Technologies, Grand Island, NY). An anti-laminin antibody (1:400, rabbit polyclonal, Z0097; Dako, Trappes, France) coupled to Alexa Fluor 488 GAR (1:200; Life Technologies) was used to delineate muscle fiber architecture. Hoechst staining was further performed to visualize nuclei. Images were visualized using a microscope (Olympus, Tokyo, Japan) and digitized using a charge-coupled device camera (Olympus). TABLE. Summary of Human Muscle Biopsies Sample CTR1 CTR2 CTR3 CTR4 CTR5 CTR6 CTR7 CTR8 CTR9 CTR10 CTR11 DMD1 DMD2 DMD3 DMD4 DMD5 DMD6 DMD7
Donor Muscle Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral Quadriceps Paravertebral Quadriceps Quadriceps Quadriceps Fascia lata Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral
Age, years 19 17 18 15 17 14 15 10 19 37 33 16 15 12 15 12 15 12
Sex M M M F M F F F M M F M M M M M M M
DMD8 DMD9 DMD10
Paravertebral Paravertebral Fascia lata
12 15 8
M M M
Mutation None None None None None None None None None None None $10Y41 c.IVS66+1G9A $3Y9 $50Y52 $8Y43 c.10141C9T, p.Arg3381X c.2281_2285delGAAAA; p.Glu761SerfsX10 c.6746dupA ND ND
$, base pair segment deletion; F, female; M, male; ND, not determined.
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GAG Extraction and Quantification At least 30 mg of frozen DMD and healthy muscle biopsies was freeze-dried and suspended in an extraction buffer for further GAG extraction, as previously described (24). Briefly, samples were digested with proteinase K (Merck, Darmstadt, Germany) and DNase (Qiagen, Courtabouef, France) and then transferred to a 4 mol/L NaCl solution. Proteins were precipitated and supernatants were cleared by chloroform washing, then the aqueous phase (Spectrum Laboratories, Breda, the Netherlands) was dialyzed against the extraction buffer and then pure water. After freeze-drying, the identities of extracted GAGs were determined by specific digestion with chondroitinase ABC (Sigma-Aldrich) or by nitrous acid treatment and quantified according to the 1Y9 dimethyl-methylene blue assay based on sulfate group complexation, as previously described (28).
Agarose Gel Electrophoresis of GAGs
About 1 to 2 Kg of GAGs isolated from CTR and DMD biopsies was separated on a 2% agarose gel run in TAE 1 buffer for 150 minutes at 50 mV. After migration, the gel was stained with 0.005% Stains-All solution (Sigma-Aldrich) overnight in the dark and destained successively with water.
CS Disaccharides Analysis by High-Performance Liquid Chromatography Disaccharides of purified CS were obtained by treatment with chondroitinase ABC (20 mU, 90 minutes, 37-C). Samples were analyzed by high-performance liquid chromatography, as previously described (29). Briefly, 50 KL (40 Kg/mL) of sample was loaded onto a Propac PA-1 (Dionex) strong anion exchange column eluted by a NaCl-solvent gradient. Postcolumn in-line modification was performed by mixing 2-cyanoacetamide solution (2% vol/vol) and 250 mmol/L NaOH, both supplied at 0.25 mL/minute. The mixture passed through a reaction coil set in an oven at 120-C then through a cooling coil, and then fluorimetric monitoring was monitored (L = 346 nm excitation; L = 410 nm emission). Areas under the curve were measured, and the percentage of each disaccharide in each sample was calculated relative to external standards.
Reverse TranscriptionYPolymerase Chain Reaction and Real-Time Reverse TranscriptionYPolymerase Chain Reaction Analysis Total RNA was extracted from tissues with Trizol reagent (Invitrogen/Life Technologies, Saint Aubin, France) and treated with RNase-Free DNase (Qiagen). Subsequently, 400 ng of RNA was reverse-transcribed using M-MuLV First Strand kit (Invitrogen), according to the manufacturer’s instructions. Quantitative polymerase chain reaction (PCR) was carried out using SYBR green mix buffer (Roche Applied Science, Meylan, France) in a LightCycler 480 Real-Time PCR System (Roche Applied Science) as follows: 8 minutes at 95-C followed by 50 cycles at 95-C for 15 seconds, 60-C for 15 seconds and 72-C for 15 seconds, with the program ending in 5 seconds at 95-C and 1 minute at 65-C. Specificity of the PCR product was checked by melting curve analysis using the following program: 65-C increasing by 0.11-C/
Chondroitin Sulfate Remodeling in DMD
second to 97-C. The expression level of each messenger RNA (mRNA) was normalized to that of B2M mRNA expression. For carbohydrate sulfotransferase (CHST) 13/C4ST-3 and CS hydrolase-4 (HYAL-4/CSHY), reverse transcription (RT) PCR was performed using a 2 ReddyMix PCR master mix (Thermo Scientific, Rockford, IL) on a thermocycler GeneAmp PCR System 2700 (Applied Biosystems, Saint Aubin, France) as follows: 30 seconds at 95-C, 30 seconds at 60-C, 30 seconds at 72-C (36 cycles for CHST13 and HYAL-4; 30 cycles for B2M). Amplification products were separated on a 2% agarose gel containing ethidium bromide for visualization, and quantification was performed using Gel Doc 2000 software (BioRad). Primer sequences used in this study are as indicated for transforming growth factor-A (TGFA), 5¶-CGCG TGCTAATGGTGGAAAC-3¶ and 5¶-GTTCAGGTACCGC TTCTCGG-3¶; for CTHRC1, 5¶-GGACCAAGGAAGCCCT GAAAT-3¶ and 5¶-AGCAACATCCACTAATCCAGCA-3¶; for osteopontin (OPN), 5¶-GCCGAGGTGATAGTGTGGTT-3¶ and 5¶-AACGGGGATGGCCTTGTATG-3¶; for CHSY1, 5¶GAAGGTGTGTCCGGAGGTTT-3¶ and 5¶-CCCCTTTTTGT TCTGCTCGT-3¶; for CHPF, 5¶-CACATGTACCAGCTGCAC AAA-3¶ and 5¶-GGCTGGTATTCTGGATCTCCC-3¶; for UST, 5¶-CTTCAAGGGCGTGCTCAGTA-3¶ and 5¶-GGGGA CAGTCTTCTTCACCG-3¶; for DSE, 5¶-TCGTCCAGAGGC ACTTCAAC-3¶ and 5¶-AGTCCGCAATAGCCACAGTC-3¶; for CHST3, 5¶-GTGCACAGCCTGAAGATGAG-3¶ and 5¶-TG TTGGCATCTGCTAGAGCTT-3¶; for CHST7, 5¶-TCTATG GCAGGCGCTGTATC-3¶ and 5¶-TGACCTTGTTAGTCCG CCAG-3¶; for CHST11, 5¶-GAGGAATCCCTTTGGTGTGG3¶ and 5¶-AGGACAGCAGTGTTTGAGAGC-3¶; for CHST12, 5¶-GAGGACTGGTTCGCCAAGAT-3¶ and 5¶-AGAGAACA AAGTCGGCCTCG-3¶; for CHST13, 5¶-TTTTCAACTACTC CGCCCCC-3¶ and 5¶-GGTCTTGTCGGAAAGGCACT-3¶; for CHST14, 5¶-AAGTTCCTGTTTGTGCGGGA-3¶ and 5¶-CCC ATAGCGTTGCTGGTACT-3¶; for CHST15, 5¶-CTCTGCAA AGGAGCAGAGCA-3¶ and 5¶-CCATCCGTGCTGTTGTCG TA-3¶; for HYAL-4, 5¶-CGCTTCTCCAAATTTCGGGTG-3¶ and 5¶-CTCCCAAGGCAGCACTTTCT-3¶; for B2M, 5¶TTTCTGGCCTGGAGGCTATC-3¶ and 5¶-TCCATTCTCTG CTGGATGACG-3¶.
Statistical Analysis All data are expressed as mean T SEM. Student t-test was used to compare differences between CTR and DMD biopsies using GraphPad Prism. A difference was considered to be significant at * p G 0.05, ** p G 0.01, or *** p G 0.001.
RESULTS Characterization of DMD Muscle Biopsies Hematoxylin and eosin staining of CTR muscles showed an organized architecture with myofibers characterized by a polygonal shape and peripheral nuclei (Fig. 1A). Biopsies of DMD patients showed a marked loss of muscle tissue accompanied by infiltration of adipose and fibrotic tissues. Residual fibers showed high variability in size and the presence of internal myonuclei (Fig. 1B). Immunostaining with anti-NNmyoHC (a marker of ongoing muscle regeneration) revealed marked degeneration/regeneration in DMD muscle
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FIGURE 1. Characteristics of human skeletal muscle biopsies. (A, B) Hematoxylin and eosin staining of control (CTR) (A) and DMD patient (B) biopsies. (C, D) Representative transverse cryosections of CTR (C) and DMD (D) biopsies stained with antibodies against NNmyoHC (red) and spectrin (green). Nuclei are counterstained with Hoechst (blue). (E) Quantification of the number of NNmyoHC-positive fibers in CTR and DMD biopsies. (F, G) Sirius red staining of CTR (F) and DMD (G) biopsies. (H) Quantification of Sirius red staining of fibrotic tissue in CTR and DMD biopsies. (IYK) Transforming growth factor-A (I), OPN (J), collagen triple helix repeat containing 1 (CTHCR1) (K) mRNA expression in CTR and DMD muscle as measured by quantitative RT-PCR. All data are expressed as mean T SEM. * p G 0.05, ** p G 0.01, *** p G 0.001. Scale bar = 50 Km.
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FIGURE 2. Distribution of HS, CS, and DS in skeletal muscle biopsies. Staining of skeletal muscle cryosections from control (CTR) (A, B, E, F, I, J) and DMD patient (C, D, G, H, K, L) biopsies with antibodies to various GAGs. Cryosections were incubated with antiHS antibody (AO4B08) (AYD), anti-CS antibody (IO3H1O) (EYH), and anti-DS antibody (LKN1) (IYL) (all in red). Laminin immunostaining (green) was added to delineate the muscle structure. Merged images are shown in B, D, F, H, J, and L. Nuclei are counterstained with Hoechst. Scale bar = 100 Km.
biopsies, which was totally absent in CTR muscles (Figs. 1CYE). Sirius red staining of CTR muscles revealed a minimal amount of connective tissue (5.4% T 0.3%) (Figs. 1F, H), whereas Sirius red staining of DMD biopsies showed a large accumulation of fibrotic tissue in the endomysial space, occupying approximately 30% of the total area (29.4% T 3.2%; *** p G 0.001) (Figs. 1G, H). Consistent with an advanced stage of the disease, no evidence of inflammation, as detected by immunostaining for CD4, CD3, and CD8, was found in the DMD biopsies (data not shown). For further characterization of fibrosis in DMD muscle biopsies, the expression of TGFA, OPN, and CTHCR1 mRNAs, which are classically upregulated in fibrotic processes, was measured by quantitative RT-PCR. All 3 mRNAs showed upregulation in DMD biopsies (Figs. 1IYK). Overall, these results confirm the presence of extensive fibrosis in DMD muscle biopsies.
GAG Expression and Distribution To investigate the expression and distribution of different GAGs in skeletal muscle, we performed immunostaining of CTR and DMD muscle biopsies, using antibodies against the sulfated GAGs expressed in skeletal muscle in combination with an anti-laminin antibody, to delineate muscle basal lamina (Fig. 2). Glycosaminoglycans were detected using 3 phage display antibodies. No difference in distribu-
tion between CTR and DMD was observed with the antibody against HS: in both biopsies, HS staining was of similar intensity and colocalized with the basal lamina (Figs. 2AYD). In contrast, immunostaining with anti-CS and anti-DS antibodies showed a very strong and diffuse staining in the ECM in DMD versus CTR muscle, suggesting large accumulation of CS and DS in the dystrophic environment (Figs. 2EYL).
Selective Accumulation of CS/DS in DMD Muscle Biopsies Quantification of total sulfated GAGs showed a significant increase in the total amount of GAGs in DMD biopsies (1.10 T 0.10 Kg/mg) versus CTR biopsies (0.79 T 0.07 Kg/mg) (p G 0.05) (Fig. 3A), suggesting increased production of ECM during the fibrotic process that accompanies the loss of muscle fibers. Specific quantification of each GAG species showed that the increase in the total amount of sulfated GAGs in DMD muscle biopsies was attributable to a selective and significant increase in CS/DS (0.73 T 0.08 Kg/mg) versus CTR (0.52 T 0.04 Kg/mg) (p G 0.01), whereas the amounts of HS did not differ between CTR and DMD biopsies (CTR, 0.43 T 0.05 Kg/mg; DMD, 0.36 T 0.03 Kg/mg) (Fig. 3A). These results demonstrate for the first time that levels of CS are specifically increased after tissue remodeling during the pathologic process in DMD patients.
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T 3.5% vs 53.0% T 7.5%) (* p G 0.05) whereas the proportion of monosulfated (MonoS) disaccharides was greater (59.0% T 3.6% vs 43.0% T 6.9%), although the latter comparison was not significant (p = 0.0608). No changes in the relative composition of disulfated disaccharides in DMD versus CTR biopsies (5.0% T 0.4% vs 4.0% T 0.6%) (Fig. 3C) were observed. These results indicate that the increase in CS/DS is not attributable to a longer chain length but to a higher proportion of MonoS disaccharide units in the DMD biopsies.
CS/DS Metabolic Enzyme Expression
FIGURE 3. Quantification and structural characterization of skeletal muscle GAGs. (A) Total sulfated GAG content in skeletal muscle from control (CTR; white histograms) and DMD patient (gray histograms) biopsies quantified using the 1Y9 dimethyl-methylene blue assay. CS and HS were quantified after nitrous acid treatment or digestion with chondroitinase ABC, respectively, using the same assay. (B) Length of GAG chains visualized by agarose gel electrophoresis. Three different CTR (1Y3) and DMD (4Y6) samples were loaded. Commercial hyaluronic acid was used as a molecular weight (MW) marker. (C) Disaccharide composition of CS in CTR (white histograms) and DMD (gray histograms) skeletal muscle biopsies determined by high-performance liquid chromatography analysis. All data are expressed as mean T SEM. * p G 0.05, ** p G 0.01. DiS, disulfated; NoS, nonsulfated; MonoS, monosulfated.
To characterize the mechanisms involved in the increased accumulation and modified sulfation of CS/DS in skeletal muscle biopsies from DMD patients versus CTR biopsies, we analyzed the expression of enzymes involved in CS/DS biosynthesis by quantitative and semiquantitative RTPCR (Fig. 4). CS/DS are composed of repeated disaccharides units of GlcA or IdoA and N-acetyl-galactosamine (GalNAc). The expression of both CS synthase CHSY1/CSS-1 and CHPF/CSS-2, which catalyze the polymerization of CS/DS, did not differ between CTR and DMD biopsies (Figs. 4A, B); this is consistent with the result in Figure 3C, which indicates that the length of the CS/DS chains was not modified in DMD. In addition, the levels of enzymes involved in GlcA/ IdoA modifications, UST (uronyl-2-sulfotransferase) and DSE (dermatan sulfate epimerase) were not different between CTR and DMD (Figs. 4C, D). Among the 3 different isoforms of the C4ST enzyme (CHST11/C4ST-1, CHST12/C4ST-2, and CHST13/C4ST-3), which catalyze the 4-O-sulfation of GalNAc (Figs. 4EYG), only CHST11 was significantly greater in DMD versus CTR biopsies (* p G 0.05) (Fig. 4E). This is in accordance with the increased amount of MonoS disaccharides observed in the DMD muscle biopsies (Fig. 3C). The expression of CHST14/D4ST-1 enzyme (which is responsible for the 4-O-sulfation of GalNAc adjacent to IdoA), CHST3/C6ST-1 and CHST7/C6ST-2 enzymes (which catalyze the 6-O-sulfation of GalNAc), and GalNAc4S-6ST/CHST15 enzyme (which transfers a sulfate group on GalNAc-4S) did not differ between DMD and CTR biopsies (Figs. 4HYK). Interestingly, the expression of the CS hydrolase HYAL-4/ CSHY enzyme, which is involved in the depolymerization of CS, was downregulated in DMD samples (** p G 0.01), suggesting diminished degradation of CS/DS in the skeletal muscle of DMD patients (Fig. 4L). All of these results are consistent with an increased level of sulfation caused by an increased proportion of MonoS disaccharides and a decreased turnover of CS/DS.
CS/DS Structural Modifications Because increased amounts of CS/DS, as measured by the 1Y9 dimethyl-methylene blue assay, may be attributable to an increase in GAG chain length and/or an increase in disaccharide unit sulfation, we assessed GAG chain length using agarose gel electrophoresis. We observed no difference in GAG chain lengths extracted from CTR and DMD biopsies (Fig. 3B). High-performance liquid chromatography analysis, however, allows one to define the relative proportion of each of the disaccharide units in the composition of CS/DS chains. In DMD patient biopsies, the proportion of nonsulfated disaccharides was significantly less than in CTR biopsies (36.0%
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DISCUSSION Extracellular matrix remodeling and fibrosis represent key components of the pathologic evolution in DMD. Although innovative therapeutic strategies are being tested to restore dystrophin expression (such as exon-skipping, gene therapy, or cell therapy), none of these will target the fibrosis that may limit access of the therapeutic agent to the muscle fibers. Fibrosis is a complex but not yet fully understood process characterized by excessive accumulation of collagens and ECM components (6). Recently, it was demonstrated in DMD that endomysial fibrosis was the only myopathologic parameter Ó 2014 American Association of Neuropathologists, Inc.
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FIGURE 4. Analysis of enzymes involved in CS/DS metabolism. Analysis of enzymes involved in CS/DS metabolism in control (CTR) and DMD patient skeletal muscle biopsies by real-time RT-PCR (AYF, HYK) and RT-PCR (G, L). Values were normalized using B2M mRNA as reference. Each value is presented as mean T SEM. Student t-test was used to compare differences between CTR and DMD biopsies. Differences were considered significant at * p G 0.05 or ** p G 0.01. CHPF, chondroitin polymerizing factor; CHSY1, CS synthase-1; DSE, DS epimerase; GlcA, glucuronic acid; IdoA, iduronic acid; UST, uronyl-2-sulfotransferase.
that significantly correlated with poor motor outcome (30). Studying matrix components could help to understand mechanisms involved in the deposition of fibrosis and to find therapeutic targets to reverse fibrosis.
CS/DS, But Not HS, Are Increased in DMD This is the first study to investigate GAG content in DMD. The most striking and original finding in our study is the marked and selective increase in CS/DS in the muscles of DMD patients, whereas no changes in the amount of HS were observed between CTR and DMD biopsies; these results were found by both histochemical and biochemical quantification of GAGs. Advanced stages of DMD are characterized by an absence of muscle regeneration caused by a depletion of the satellite cell compartment in chronically damaged DMD muscle. To date, many studies have been conducted on syndecans, a family of PGs that carry both HS and CS chains, highlighting their roles as regulators of skeletal muscle regeneration, probably through modulation of signaling pathways by HS chains (17, 31, 32). It is possible that at this stage of the disease, HSmediated mechanisms, which participate in the regulation of muscle regeneration, are no longer active. The histochemical
approach localized CS both in the body of the ECM and in the basal lamina, whereas DS was in the endomysial space of the skeletal muscle. These results are in agreement with previous histochemical studies on PGs, which demonstrated that CS/DSPG biglycan and DS-PG decorin accumulate throughout the perimysium and endomysium (33). Recently, a study using a similar immunostaining approach performed on human skeletal muscle showed a decrease in HS and an increase in CS in the diaphragm muscles of patients affected by chronic obstructive pulmonary disease, which is associated with dysfunction of inspiratory muscles (34). Altogether, these results indicate that a selective increase in CS/DS content represents a specific response to a controlled mechanism that induces ECM remodeling in dystrophic and other pathologically altered muscles.
DMD Is Characterized by an Increased Proportion of CS/DS 4-O-Sulfation In addition to the accumulation of CS/DS in DMD, we demonstrate for the first time that GAG fine structure is also modified as well. We observed an enriched proportion of MonoS units and probably a specific increase in 4-O-sulfation of GalNAc residues, consistent with the upregulation of
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the CHST11/C4ST-1 enzyme. Our results are in agreement with a previous study, which suggested that the CHST11/ C4ST-1 isoform acts in cooperation with the chondroitin Nacetylgalactosaminyltransferase 2 to increase the amount of CS biosynthesized by cells (35). We also found that the expression of the CS hydrolase HYAL-4/CSHY enzyme, which is involved in the depolymerization of CS (36Y38), was downregulated in DMD samples. Because we found no modifications in GAG chain length in DMD, the increase in CS/DS in DMD biopsies can also be explained by an increase in the number of GAG chains, consistent with a diminished activity of HYAL-4/CSHY. Together, these results suggest that, in DMD patients, CS chains become increasingly sulfated and their turnover is reduced. Interestingly, analysis of CS/DS metabolic enzymes in the affected cricopharyngeal muscle of oculopharyngeal muscular dystrophy patients, where atrophy and fibrosis are present (39), showed a different regulation of the machinery of sulfation of CS/DS (data not shown). This indicates that the modifications in GAG sulfation observed in DMD are not a general muscle fibrotic process and could be specific for DMD. Recently, Mikami et al (40) showed that levels of CS chains in the murine myoblast cell line C2C12 are dramatically diminished at the stage of extensive syncytial myotube formation, indicating that a decrease in CS content, mediated by Hyal-1 activity, is required for muscle differentiation. In addition, they showed that enzymatic removal of all CS species improved the dystrophic pathology in the muscles of mdx mice, a model for DMD. Therefore, it would be of interest to determine if the upregulation of CS/DS and, more specifically, the increasing proportion of 4-O-sulfation in human dystrophic muscle are associated with changes in GAG functions. We have previously demonstrated that GAGs from an aged myocardium present altered sulfation patterns, leading to disruption of growth factor binding and activities (24). We showed that GAGs from aged cardiac muscle display better affinity for vascular endothelial growth factor, whereas aging significantly reduces the capacity of these GAGs to bind fibroblast growth factor-1 and fibroblast growth factor-2. Interestingly, TGFA (one of the key players in muscle tissue remodeling) (6), CTGF (41), and OPN (42) are all upregulated in DMD (41Y43), and each of them is a HBPs, which is regulated by GAG chains. In addition, growth factors of the TGFA superfamily have been identified as positive regulators of the expression of the CHST11/C4ST-1 gene (44), suggesting a possible positive feedback loop on fibrosis. In conclusion, we describe for the first time an accumulation of CS/DS GAGs in DMD muscle and a modification of the sulfation pattern of CS/DS in DMD biopsies, suggesting a possible role for 2 enzymes (CHST11/C4ST-1 and HYAL-4/CSHY) implicated in the machinery of sulfation of CS/DS. Understanding of the roles of GAGs in muscular dystrophies has only started to emerge. Thus, it remains to be determined if the alteration of GAGs is a consequence of the primary remodeling that contributes to worsening of the disease and then participates in the formation of fibrosis or, in contrast, if their modulation is a compensatory process. In DMD, further studies using new glycomic technologies are required to more finely characterize CS/DS functionality in
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muscle biopsies to determine the relationship between the changes occurring in CS/DS sulfation pattern and their ability to control such HBP-induced signaling, leading to the formation and maintenance of fibrosis. Finally, GAG fine structure characterization in DMD biopsies may provide a new tool for the design of a potential new class of anti-fibrotic drug candidates. ACKNOWLEDGMENTS We gratefully thank C. Trollet, A. Bigot, and P. Klein for fruitful discussions and critical reading of the manuscript. We also thank G. Butler-Browne and V. Mouly for their support and for precious discussions during the preparation of the manuscript. We thank S. Vasseur and M. Chapart from Myobank for providing muscle biopsies. We thank OTR3 (Paris, France) for assuring M. Gilbert’s salary independently of this work. REFERENCES 1. Mendell JR, Shilling C, Leslie ND, et al. Evidence-based path to newborn screening for Duchenne muscular dystrophy. Ann Neurol 2012;71:304Y13 2. Hoffman EP, Brown RH Jr, Kunkel LM. Dystrophin: The protein product of the Duchenne muscular dystrophy locus. Cell 1987;51:919Y28 3. Emery AE. The muscular dystrophies. Lancet 2002;359:687Y95 4. Blake DJ, Weir A, Newey SE, et al. Function and genetics of dystrophin and dystrophin-related proteins in muscle. Physiol Rev 2002;82:291Y329 5. Durbeej M, Campbell KP. Muscular dystrophies involving the dystrophinglycoprotein complex: An overview of current mouse models. Curr Opin Genet Dev 2002;12:349Y61 6. Serrano AL, Munoz-Canoves P. Regulation and dysregulation of fibrosis in skeletal muscle. Exp Cell Res 2010;316:3050Y58 7. Gargioli C, Coletta M, De Grandis F, et al. PlGF-MMP-9Yexpressing cells restore microcirculation and efficacy of cell therapy in aged dystrophic muscle. Nat Med 2008;14:973Y78 8. Le Hir M, Goyenvalle A, Peccate C, et al. AAV genome loss from dystrophic mouse muscles during AAV-U7 snRNA-mediated exon-skipping therapy. Mol Ther 2013;21:1551Y58 9. Raghow R. The role of extracellular matrix in postinflammatory wound healing and fibrosis. FASEB J 1994;8:823Y31 10. Frantz C, Stewart KM, Weaver VM. The extracellular matrix at a glance. J Cell Sci 2010;123:4195Y200 11. Sugahara K, Mikami T, Uyama T, et al. Recent advances in the structural biology of chondroitin sulfate and dermatan sulfate. Curr Opin Struct Biol 2003;13:612Y20 12. Volpi N. Therapeutic applications of glycosaminoglycans. Curr Med Chem 2006;13:1799Y810 13. Jenniskens GJ, Veerkamp JH, van Kuppevelt TH. Heparan sulfates in skeletal muscle development and physiology. J Cell Physiol 2006;206: 283Y94 14. Zhang L. Glycosaminoglycan (GAG) biosynthesis and GAG-binding proteins. Prog Mol Biol Transl Sci 2010;93:1Y17 15. Handel TM, Johnson Z, Crown SE, et al. Regulation of protein function by glycosaminoglycansVas exemplified by chemokines. Annu Rev Biochem 2005;74:385Y410 16. Friedl A, Chang Z, Tierney A, et al. Differential binding of fibroblast growth factor-2 and -7 to basement membrane heparan sulfate: Comparison of normal and abnormal human tissues. Am J Pathol 1997;150: 1443Y55 17. Pisconti A, Bernet JD, Olwin BB. Syndecans in skeletal muscle development, regeneration and homeostasis. Muscles Ligaments Tendons J 2012;2:1Y9 18. Caceres S, Cuellar C, Casar JC, et al. Synthesis of proteoglycans is augmented in dystrophic mdx mouse skeletal muscle. Eur J Cell Biol 2000;79:173Y81 19. Casar JC, McKechnie BA, Fallon JR, et al. Transient up-regulation of biglycan during skeletal muscle regeneration: Delayed fiber growth along with decorin increase in biglycan-deficient mice. Dev Biol 2004;268: 358Y71
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J Neuropathol Exp Neurol Volume 73, Number 8, August 2014 20. Haslett JN, Kunkel LM. Microarray analysis of normal and dystrophic skeletal muscle. Int J Dev Neurosci 2002;20:359Y65 21. Zanotti S, Negri T, Cappelletti C, et al. Decorin and biglycan expression is differentially altered in several muscular dystrophies. Brain 2005;128: 2546Y55 22. Riederer I, Negroni E, Bencze M, et al. Slowing down differentiation of engrafted human myoblasts into immunodeficient mice correlates with increased proliferation and migration. Mol Ther 2012;20:146Y54 23. Butler-Browne GS, Whalen RG. Myosin isozyme transitions occurring during the postnatal development of the rat soleus muscle. Dev Biol 1984; 102:324Y34 24. Huynh MB, Morin C, Carpentier G, et al. Age-related changes in rat myocardium involve altered capacities of glycosaminoglycans to potentiate growth factor functions and heparan sulfateYaltered sulfation. J Biol Chem 2012;287:11363Y73 25. Kurup S, Wijnhoven TJ, Jenniskens GJ, et al. Characterization of antiYheparan sulfate phage display antibodies AO4B08 and HS4E4. J Biol Chem 2007;282:21032Y42 26. Smetsers TF, van de Westerlo EM, ten Dam GB, et al. Human single-chain antibodies reactive with native chondroitin sulfate detect chondroitin sulfate alterations in melanoma and psoriasis. J Invest Dermatol 2004;122:707Y16 27. Lensen JF, Wijnhoven TJ, Kuik LH, et al. Selection and characterization of a unique phage displayYderived antibody against dermatan sulfate. Matrix Biol 2006;25:457Y61 28. Barbosa I, Garcia S, Barbier-Chassefiere V, et al. Improved and simple micro assay for sulfated glycosaminoglycans quantification in biological extracts and its use in skin and muscle tissue studies. Glycobiology 2003; 13:647Y53 29. Toyoda H, Yamamoto H, Ogino N, et al. Rapid and sensitive analysis of disaccharide composition in heparin and heparan sulfate by reversed-phase ion-pair chromatography on a 2 Km porous silica gel column. J Chromatogr A 1999;830:197Y201 30. Desguerre I, Christov C, Mayer M, et al. Clinical heterogeneity of Duchenne muscular dystrophy (DMD): Definition of sub-phenotypes and predictive criteria by long-term follow-up. PLoS One 2009;4:e4347 31. Cornelison DD, Filla MS, Stanley HM, et al. Syndecan-3 and syndecan-4 specifically mark skeletal muscle satellite cells and are implicated in satellite cell maintenance and muscle regeneration. Dev Biol 2001;239:79Y94
Chondroitin Sulfate Remodeling in DMD 32. Cornelison DD, Wilcox-Adelman SA, Goetinck PF, et al. Essential and separable roles for Syndecan-3 and Syndecan-4 in skeletal muscle development and regeneration. Genes Dev 2004;18:2231Y36 33. Carrino DA, Caplan AI. Isolation and preliminary characterization of proteoglycans synthesized by skeletal muscle. J Biol Chem 1982;257: 14145Y54 34. Ottenheijm CA, Jenniskens GJ, Geraedts MC, et al. Diaphragm dysfunction in chronic obstructive pulmonary disease: A role for heparan sulphate? Eur Respir J 2007;30:80Y89 35. Izumikawa T, Okuura Y, Koike T, et al. Chondroitin 4-O-sulfotransferase-1 regulates the chain length of chondroitin sulfate in co-operation with chondroitin N-acetylgalactosaminyltransferase-2. Biochem J 2011;434: 321Y31 36. Csoka AB, Frost GI, Stern R. The six hyaluronidase-like genes in the human and mouse genomes. Matrix Biol 2001;20:499Y508 37. Stern R, Jedrzejas MJ. Hyaluronidases. their genomics, structures, and mechanisms of action. Chem Rev 2006;106:818Y39 38. Kaneiwa T, Mizumoto S, Sugahara K, et al. Identification of human hyaluronidase-4 as a novel chondroitin sulfate hydrolase that preferentially cleaves the galactosaminidic linkage in the trisulfated tetrasaccharide sequence. Glycobiology 2010;20:300Y9 39. Gidaro T, Negroni E, Perie´ S, et al. Atrophy, fibrosis and PAX7-positive cells in pharyngeal muscles of oculopharyngeal muscular dystrophy patients. J Neuropathol Exp Neurol 2013;72(3):234Y43 40. Mikami T, Koyama S, Yabuta Y, et al. Chondroitin sulfate is a crucial determinant for skeletal muscle development/regeneration and improvement of muscular dystrophies. J Biol Chem 2012;287:38531Y42 41. Sun G, Haginoya K, Wu Y, et al. Connective tissue growth factor is overexpressed in muscles of human muscular dystrophy. J Neurol Sci 2008;267:48Y56 42. Vetrone SA, Montecino-Rodriguez E, Kudryashova E, et al. Osteopontin promotes fibrosis in dystrophic mouse muscle by modulating immune cell subsets and intramuscular TGF-beta. J Clin Invest 2009;119:1583Y94 43. Bernasconi P, Di Blasi C, Mora M, et al. Transforming growth factor-beta1 and fibrosis in congenital muscular dystrophies. Neuromuscul Disord 1999;9:28Y33 44. Willis CM, Wrana JL, Kluppel M. Identification and characterization of TGFbeta-dependent and -independent cis-regulatory modules in the C4ST1/CHST11 locus. Genet Mol Res 2009;8:1331Y43
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ORIGINAL ARTICLE
Glycosaminoglycan Modifications in Duchenne Muscular Dystrophy: Specific Remodeling of Chondroitin Sulfate/ Dermatan Sulfate Elisa Negroni, PhD, Emilie Henault, MSc, Fabien Chevalier, PhD, Marie Gilbert-Sirieix, PhD, Toin H. Van Kuppevelt, PhD, Dulce Papy-Garcia, PhD, Georges Uzan, PhD, and Patricia Albanese, PhD
Abstract Widespread skeletal muscle degeneration and impaired regeneration lead to progressive muscle weakness and premature death in patients with Duchenne muscular dystrophy (DMD). Dystrophic muscles are progressively replaced by nonfunctional tissue because of exhaustion of muscle precursor cells and excessive accumulation of extracellular matrix (ECM). Sulfated glycosaminoglycans (GAGs) are components of the ECM and are increasingly implicated in the regulation of biologic processes, but their possible role in the progression of DMD pathology is not understood. In the present study, we performed immunohistochemical and biochemical analyses of endogenous GAGs in skeletal muscle biopsies of 10 DMD patients and 11 healthy individuals (controls). Immunostaining targeted to specific GAG species showed greater deposition of chondroitin sulfate (CS)/dermatan (DS) sulfate in DMD patient biopsies versus control biopsies. The selective accumulation of CS/DS in DMD biopsies was confirmed by biochemical quantification assay. In addition, highperformance liquid chromatography analysis demonstrated a modification of the sulfation pattern of CS/DS disaccharide units in DMD muscles. In conclusion, our data open up a new path of investigation and suggest that GAGs could represent a new and original therapeutic target for improving the success of gene or cell therapy for the treatment of muscular dystrophies.
From the INSERM U972, Paul Brousse Hospital, Villejuif (EN, FC, GU); and EAC CNRS 7149, CRRET Laboratory, Sciences and Technology Faculty, Paris-Est Creteil University, Cre´teil (EH, FC, MG-S, DP-G, PA), France; and Department of Biochemistry (280), Nijmegen Center for Molecular Life Sciences, Radboud University Nijmegen Medical Center, Nijmegen, the Netherlands (THVK). Send correspondence and reprint requests to: Elisa Negroni, PhD, INSERM U972, Paul Brousse Hospital, 12 Avenue Paul Vaillant Couturier, Villejuif 94807, France; E-mail: [email protected] Present address: Elisa Negroni, UMRS 974, INSERM, Universite´ Paris 6, CNRS UMR 7215, Institut de Myologie, Baˆtiment Babinski, Groupe Hospitalier Pitie´-Salpeˆtrie`re 47, boulevard de l’Hoˆpital, 75651 Paris Cedex 13, France. Georges Uzan and Patricia Albanese contributed equally to this work. This study was supported by grants from Association Fran0aise contre les Myopathies and Region Ile-de-France. Elisa Negroni was supported by a fellowship from Association Fran0aise contre les Myopathies. Fabien Chevalier was supported by a doctoral fellowship from Region Ile-de-France. All authors declare no conflicts of interest.
Key Words: Chondroitin sulfate, Dermatan sulfate, Duchenne muscular dystrophy, Extracellular matrix, Fibrosis, Glycosaminoglycans.
INTRODUCTION Duchenne muscular dystrophy (DMD) is the most common inherited muscular dystrophy of childhood, affecting 1 in 5,000 live male births (1). It is caused by mutations in the dystrophin gene, leading to the absence of a functional dystrophin protein (2). In DMD patients, widespread skeletal muscle degeneration most often leads to progressive muscle wasting, cardiac dysfunction, respiratory failure, and, ultimately, death in the third decade of life (3). Dystrophin links the extracellular matrix (ECM) to the cytoskeleton of muscle fibers through the dystrophin-associated glycoprotein complex (4); it is a cytoskeletal protein essential for the structural integrity of the membrane of myofibers (i.e. the sarcolemma) (5). Absence of dystrophin results in permanent fragility and leakiness of the sarcolemma, which lead to increased calcium influx and progressive degeneration and necrosis of muscle fibers with subsequent loss of muscle tissue and progressive fibrosis (6). Fibrosis appears early in muscles of DMD patients, increases with age, and represents a major hurdle hampering the success of gene or cell therapies at advanced stages of the disease (7, 8). Indeed, fibrosis is a complex process characterized by excessive accumulation of ECM components in which there is progressive replacement of functional muscle tissue by nonfunctional connective tissue, leading to worsening of the disease and irreversible loss of muscle function. In normal tissue, the ECM is a dynamic structure that has important roles in cell signaling and cell anchorage, thus maintaining tissue homeostasis (9). It is mainly composed of structural glycoproteins and fibrous proteins such as collagens and proteoglycans (PGs) (10). Proteoglycans are the most abundant components of the nonfibrillar ECM. They are composed of a protein core to which long, linear, highly sulfated glycosaminoglycan (GAG) chains are covalently at tached. Proteoglycans display different functions that are principally mediated by the GAG chains (11). Glycosaminoglycans are polymers of alternating disaccharide residues. Their structure is highly complex owing to the nature of repeating disaccharide units, the heterogeneity of glycosidic linkage, sulfation patterns, and hydrophobic regions, which
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regulate their interactions with surrounding molecules (12, 13). Studying GAG signaling is complex because their biologic functions are directly related to their fine structure. Moreover, the structural variability of GAG sequences is generated during their biosynthesis under the control of multiple enzymes that progressively assemble and simultaneously modify the original polysaccharide backbone (12). The 2 main sulfated GAG families in muscle are heparan sulfate (HS) and chondroitin sulfate (CS)/dermatan sulfate (DS) (14, 15). They are involved in a variety of physiologic processes through activation, binding, or protection of various heparin-binding proteins (HBPs), such as growth factors, chemokines, cell surface receptors, and other ECM components that regulate key steps in development and tissue homeostasis (14Y16). Many studies have focused on the core proteins of PGs in skeletal muscle, irrespective of the GAG chains they harbor. For example, the role of syndecan (HS/CS-PG) core proteins has been studied in muscle development and regeneration, where they regulate satellite cell activation and myofiber formation (17). The roles of decorin (CS-PG) and biglycan (CS-PG) core proteins have been investigated in mdx mice in which these PGs have been shown to be upregulated (18, 19). In human DMD muscle, these CS-PGs appear to be altered, depending on the cohort of patients analyzed (20, 21). None of the studies describing changes in PG spatiotemporal expression during myogenesis or muscular dystrophy have focused on GAG structural epitope remodeling, which is directly related to the interaction with HBPs. In the present study, we performed immunochemical and biochemical analyses of endogenous GAGs in DMD patient and control human muscle biopsies to investigate changes in spatial distribution and structure that may contribute to the pathophysiology observed in this devastating muscle disease.
MATERIALS AND METHODS Patients Human skeletal muscle biopsies from 10 DMD patients (aged 8Y16 years) and 11 healthy individuals (controls [CTR]) were analyzed. Characteristics of the biopsies are summarized in the Table. After informed consent had been provided by the participants, all biopsies were obtained from Myobank (a partner in the European Union network EuroBioBank) in accordance with French and European Union legislation on ethical rules.
Muscle Histology Muscle biopsies were mounted on gum tragacanth (6% in water; Sigma-Aldrich, St Louis, MO) and snap-frozen in liquid N2-cooled isopentane. Staining was carried out on 5-Km-thick transverse cryosections. For the assessment of tissue morphology and visualization of fibrosis and connective tissue, muscle sections were stained with hematoxylin and eosin and Sirius red and examined by light microscopy. Fibrosis was quantified and expressed as a percentage of the surface of the whole crosssectional area on Sirius redYstained slides using Image J 1.44o analysis software (http://imagej.nih.gov/ij).
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Immunofluorescence Immunostaining was performed on 5-Km-thick cryostat muscle sections, as previously described (22). The following primary antibodies were used: antiYneonatal myosin heavy chain (NNmyoHC) (1:50, rabbit polyclonal) (23) and antispectrin (1:50, NCL-Spec1, mouse monoclonal IgG2b; Novocastra). Sections were then incubated with appropriate secondary antibodies, and nuclei were stained with Hoechst (Sigma-Aldrich). To visualize GAGs, we performed immunostaining, as previously described (24). Briefly, cryosections were fixed with 4% paraformaldehyde, incubated for 10 minutes with 50 mmol/L NH4Cl, and saturated with 3% bovine serum albumin/ phosphate-buffered saline for 45 minutes. Glycosaminoglycans were then stained overnight at 4-C with different phage display antibodies: AO4B08 (1:20), which recognizes HS species and was described to interact with an ubiquitous N-, 2-OY, and 6OYsulfated saccharide motif (25); IO3H10 antibody (1:5), which is described to recognize chondroitin 6-O-sulfate (26) and was used to reveal CS; and LKN1 antibody (1:5), which is described to recognize GlcA2S-GalNAc4S motif (27) and was used to stain DS. Immunostainings were carried out overnight at 4-C. Bound antibodies were detected with mouse antiYvesicular stomatitis virus monoclonal antibody (P5D4, 1:200) then incubated with a secondary antibody coupled to Alexa Fluor 555 GAM (1:200; Life Technologies, Grand Island, NY). An anti-laminin antibody (1:400, rabbit polyclonal, Z0097; Dako, Trappes, France) coupled to Alexa Fluor 488 GAR (1:200; Life Technologies) was used to delineate muscle fiber architecture. Hoechst staining was further performed to visualize nuclei. Images were visualized using a microscope (Olympus, Tokyo, Japan) and digitized using a charge-coupled device camera (Olympus). TABLE. Summary of Human Muscle Biopsies Sample CTR1 CTR2 CTR3 CTR4 CTR5 CTR6 CTR7 CTR8 CTR9 CTR10 CTR11 DMD1 DMD2 DMD3 DMD4 DMD5 DMD6 DMD7
Donor Muscle Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral Quadriceps Paravertebral Quadriceps Quadriceps Quadriceps Fascia lata Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral Paravertebral
Age, years 19 17 18 15 17 14 15 10 19 37 33 16 15 12 15 12 15 12
Sex M M M F M F F F M M F M M M M M M M
DMD8 DMD9 DMD10
Paravertebral Paravertebral Fascia lata
12 15 8
M M M
Mutation None None None None None None None None None None None $10Y41 c.IVS66+1G9A $3Y9 $50Y52 $8Y43 c.10141C9T, p.Arg3381X c.2281_2285delGAAAA; p.Glu761SerfsX10 c.6746dupA ND ND
$, base pair segment deletion; F, female; M, male; ND, not determined.
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GAG Extraction and Quantification At least 30 mg of frozen DMD and healthy muscle biopsies was freeze-dried and suspended in an extraction buffer for further GAG extraction, as previously described (24). Briefly, samples were digested with proteinase K (Merck, Darmstadt, Germany) and DNase (Qiagen, Courtabouef, France) and then transferred to a 4 mol/L NaCl solution. Proteins were precipitated and supernatants were cleared by chloroform washing, then the aqueous phase (Spectrum Laboratories, Breda, the Netherlands) was dialyzed against the extraction buffer and then pure water. After freeze-drying, the identities of extracted GAGs were determined by specific digestion with chondroitinase ABC (Sigma-Aldrich) or by nitrous acid treatment and quantified according to the 1Y9 dimethyl-methylene blue assay based on sulfate group complexation, as previously described (28).
Agarose Gel Electrophoresis of GAGs
About 1 to 2 Kg of GAGs isolated from CTR and DMD biopsies was separated on a 2% agarose gel run in TAE 1 buffer for 150 minutes at 50 mV. After migration, the gel was stained with 0.005% Stains-All solution (Sigma-Aldrich) overnight in the dark and destained successively with water.
CS Disaccharides Analysis by High-Performance Liquid Chromatography Disaccharides of purified CS were obtained by treatment with chondroitinase ABC (20 mU, 90 minutes, 37-C). Samples were analyzed by high-performance liquid chromatography, as previously described (29). Briefly, 50 KL (40 Kg/mL) of sample was loaded onto a Propac PA-1 (Dionex) strong anion exchange column eluted by a NaCl-solvent gradient. Postcolumn in-line modification was performed by mixing 2-cyanoacetamide solution (2% vol/vol) and 250 mmol/L NaOH, both supplied at 0.25 mL/minute. The mixture passed through a reaction coil set in an oven at 120-C then through a cooling coil, and then fluorimetric monitoring was monitored (L = 346 nm excitation; L = 410 nm emission). Areas under the curve were measured, and the percentage of each disaccharide in each sample was calculated relative to external standards.
Reverse TranscriptionYPolymerase Chain Reaction and Real-Time Reverse TranscriptionYPolymerase Chain Reaction Analysis Total RNA was extracted from tissues with Trizol reagent (Invitrogen/Life Technologies, Saint Aubin, France) and treated with RNase-Free DNase (Qiagen). Subsequently, 400 ng of RNA was reverse-transcribed using M-MuLV First Strand kit (Invitrogen), according to the manufacturer’s instructions. Quantitative polymerase chain reaction (PCR) was carried out using SYBR green mix buffer (Roche Applied Science, Meylan, France) in a LightCycler 480 Real-Time PCR System (Roche Applied Science) as follows: 8 minutes at 95-C followed by 50 cycles at 95-C for 15 seconds, 60-C for 15 seconds and 72-C for 15 seconds, with the program ending in 5 seconds at 95-C and 1 minute at 65-C. Specificity of the PCR product was checked by melting curve analysis using the following program: 65-C increasing by 0.11-C/
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second to 97-C. The expression level of each messenger RNA (mRNA) was normalized to that of B2M mRNA expression. For carbohydrate sulfotransferase (CHST) 13/C4ST-3 and CS hydrolase-4 (HYAL-4/CSHY), reverse transcription (RT) PCR was performed using a 2 ReddyMix PCR master mix (Thermo Scientific, Rockford, IL) on a thermocycler GeneAmp PCR System 2700 (Applied Biosystems, Saint Aubin, France) as follows: 30 seconds at 95-C, 30 seconds at 60-C, 30 seconds at 72-C (36 cycles for CHST13 and HYAL-4; 30 cycles for B2M). Amplification products were separated on a 2% agarose gel containing ethidium bromide for visualization, and quantification was performed using Gel Doc 2000 software (BioRad). Primer sequences used in this study are as indicated for transforming growth factor-A (TGFA), 5¶-CGCG TGCTAATGGTGGAAAC-3¶ and 5¶-GTTCAGGTACCGC TTCTCGG-3¶; for CTHRC1, 5¶-GGACCAAGGAAGCCCT GAAAT-3¶ and 5¶-AGCAACATCCACTAATCCAGCA-3¶; for osteopontin (OPN), 5¶-GCCGAGGTGATAGTGTGGTT-3¶ and 5¶-AACGGGGATGGCCTTGTATG-3¶; for CHSY1, 5¶GAAGGTGTGTCCGGAGGTTT-3¶ and 5¶-CCCCTTTTTGT TCTGCTCGT-3¶; for CHPF, 5¶-CACATGTACCAGCTGCAC AAA-3¶ and 5¶-GGCTGGTATTCTGGATCTCCC-3¶; for UST, 5¶-CTTCAAGGGCGTGCTCAGTA-3¶ and 5¶-GGGGA CAGTCTTCTTCACCG-3¶; for DSE, 5¶-TCGTCCAGAGGC ACTTCAAC-3¶ and 5¶-AGTCCGCAATAGCCACAGTC-3¶; for CHST3, 5¶-GTGCACAGCCTGAAGATGAG-3¶ and 5¶-TG TTGGCATCTGCTAGAGCTT-3¶; for CHST7, 5¶-TCTATG GCAGGCGCTGTATC-3¶ and 5¶-TGACCTTGTTAGTCCG CCAG-3¶; for CHST11, 5¶-GAGGAATCCCTTTGGTGTGG3¶ and 5¶-AGGACAGCAGTGTTTGAGAGC-3¶; for CHST12, 5¶-GAGGACTGGTTCGCCAAGAT-3¶ and 5¶-AGAGAACA AAGTCGGCCTCG-3¶; for CHST13, 5¶-TTTTCAACTACTC CGCCCCC-3¶ and 5¶-GGTCTTGTCGGAAAGGCACT-3¶; for CHST14, 5¶-AAGTTCCTGTTTGTGCGGGA-3¶ and 5¶-CCC ATAGCGTTGCTGGTACT-3¶; for CHST15, 5¶-CTCTGCAA AGGAGCAGAGCA-3¶ and 5¶-CCATCCGTGCTGTTGTCG TA-3¶; for HYAL-4, 5¶-CGCTTCTCCAAATTTCGGGTG-3¶ and 5¶-CTCCCAAGGCAGCACTTTCT-3¶; for B2M, 5¶TTTCTGGCCTGGAGGCTATC-3¶ and 5¶-TCCATTCTCTG CTGGATGACG-3¶.
Statistical Analysis All data are expressed as mean T SEM. Student t-test was used to compare differences between CTR and DMD biopsies using GraphPad Prism. A difference was considered to be significant at * p G 0.05, ** p G 0.01, or *** p G 0.001.
RESULTS Characterization of DMD Muscle Biopsies Hematoxylin and eosin staining of CTR muscles showed an organized architecture with myofibers characterized by a polygonal shape and peripheral nuclei (Fig. 1A). Biopsies of DMD patients showed a marked loss of muscle tissue accompanied by infiltration of adipose and fibrotic tissues. Residual fibers showed high variability in size and the presence of internal myonuclei (Fig. 1B). Immunostaining with anti-NNmyoHC (a marker of ongoing muscle regeneration) revealed marked degeneration/regeneration in DMD muscle
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FIGURE 1. Characteristics of human skeletal muscle biopsies. (A, B) Hematoxylin and eosin staining of control (CTR) (A) and DMD patient (B) biopsies. (C, D) Representative transverse cryosections of CTR (C) and DMD (D) biopsies stained with antibodies against NNmyoHC (red) and spectrin (green). Nuclei are counterstained with Hoechst (blue). (E) Quantification of the number of NNmyoHC-positive fibers in CTR and DMD biopsies. (F, G) Sirius red staining of CTR (F) and DMD (G) biopsies. (H) Quantification of Sirius red staining of fibrotic tissue in CTR and DMD biopsies. (IYK) Transforming growth factor-A (I), OPN (J), collagen triple helix repeat containing 1 (CTHCR1) (K) mRNA expression in CTR and DMD muscle as measured by quantitative RT-PCR. All data are expressed as mean T SEM. * p G 0.05, ** p G 0.01, *** p G 0.001. Scale bar = 50 Km.
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Chondroitin Sulfate Remodeling in DMD
FIGURE 2. Distribution of HS, CS, and DS in skeletal muscle biopsies. Staining of skeletal muscle cryosections from control (CTR) (A, B, E, F, I, J) and DMD patient (C, D, G, H, K, L) biopsies with antibodies to various GAGs. Cryosections were incubated with antiHS antibody (AO4B08) (AYD), anti-CS antibody (IO3H1O) (EYH), and anti-DS antibody (LKN1) (IYL) (all in red). Laminin immunostaining (green) was added to delineate the muscle structure. Merged images are shown in B, D, F, H, J, and L. Nuclei are counterstained with Hoechst. Scale bar = 100 Km.
biopsies, which was totally absent in CTR muscles (Figs. 1CYE). Sirius red staining of CTR muscles revealed a minimal amount of connective tissue (5.4% T 0.3%) (Figs. 1F, H), whereas Sirius red staining of DMD biopsies showed a large accumulation of fibrotic tissue in the endomysial space, occupying approximately 30% of the total area (29.4% T 3.2%; *** p G 0.001) (Figs. 1G, H). Consistent with an advanced stage of the disease, no evidence of inflammation, as detected by immunostaining for CD4, CD3, and CD8, was found in the DMD biopsies (data not shown). For further characterization of fibrosis in DMD muscle biopsies, the expression of TGFA, OPN, and CTHCR1 mRNAs, which are classically upregulated in fibrotic processes, was measured by quantitative RT-PCR. All 3 mRNAs showed upregulation in DMD biopsies (Figs. 1IYK). Overall, these results confirm the presence of extensive fibrosis in DMD muscle biopsies.
GAG Expression and Distribution To investigate the expression and distribution of different GAGs in skeletal muscle, we performed immunostaining of CTR and DMD muscle biopsies, using antibodies against the sulfated GAGs expressed in skeletal muscle in combination with an anti-laminin antibody, to delineate muscle basal lamina (Fig. 2). Glycosaminoglycans were detected using 3 phage display antibodies. No difference in distribu-
tion between CTR and DMD was observed with the antibody against HS: in both biopsies, HS staining was of similar intensity and colocalized with the basal lamina (Figs. 2AYD). In contrast, immunostaining with anti-CS and anti-DS antibodies showed a very strong and diffuse staining in the ECM in DMD versus CTR muscle, suggesting large accumulation of CS and DS in the dystrophic environment (Figs. 2EYL).
Selective Accumulation of CS/DS in DMD Muscle Biopsies Quantification of total sulfated GAGs showed a significant increase in the total amount of GAGs in DMD biopsies (1.10 T 0.10 Kg/mg) versus CTR biopsies (0.79 T 0.07 Kg/mg) (p G 0.05) (Fig. 3A), suggesting increased production of ECM during the fibrotic process that accompanies the loss of muscle fibers. Specific quantification of each GAG species showed that the increase in the total amount of sulfated GAGs in DMD muscle biopsies was attributable to a selective and significant increase in CS/DS (0.73 T 0.08 Kg/mg) versus CTR (0.52 T 0.04 Kg/mg) (p G 0.01), whereas the amounts of HS did not differ between CTR and DMD biopsies (CTR, 0.43 T 0.05 Kg/mg; DMD, 0.36 T 0.03 Kg/mg) (Fig. 3A). These results demonstrate for the first time that levels of CS are specifically increased after tissue remodeling during the pathologic process in DMD patients.
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T 3.5% vs 53.0% T 7.5%) (* p G 0.05) whereas the proportion of monosulfated (MonoS) disaccharides was greater (59.0% T 3.6% vs 43.0% T 6.9%), although the latter comparison was not significant (p = 0.0608). No changes in the relative composition of disulfated disaccharides in DMD versus CTR biopsies (5.0% T 0.4% vs 4.0% T 0.6%) (Fig. 3C) were observed. These results indicate that the increase in CS/DS is not attributable to a longer chain length but to a higher proportion of MonoS disaccharide units in the DMD biopsies.
CS/DS Metabolic Enzyme Expression
FIGURE 3. Quantification and structural characterization of skeletal muscle GAGs. (A) Total sulfated GAG content in skeletal muscle from control (CTR; white histograms) and DMD patient (gray histograms) biopsies quantified using the 1Y9 dimethyl-methylene blue assay. CS and HS were quantified after nitrous acid treatment or digestion with chondroitinase ABC, respectively, using the same assay. (B) Length of GAG chains visualized by agarose gel electrophoresis. Three different CTR (1Y3) and DMD (4Y6) samples were loaded. Commercial hyaluronic acid was used as a molecular weight (MW) marker. (C) Disaccharide composition of CS in CTR (white histograms) and DMD (gray histograms) skeletal muscle biopsies determined by high-performance liquid chromatography analysis. All data are expressed as mean T SEM. * p G 0.05, ** p G 0.01. DiS, disulfated; NoS, nonsulfated; MonoS, monosulfated.
To characterize the mechanisms involved in the increased accumulation and modified sulfation of CS/DS in skeletal muscle biopsies from DMD patients versus CTR biopsies, we analyzed the expression of enzymes involved in CS/DS biosynthesis by quantitative and semiquantitative RTPCR (Fig. 4). CS/DS are composed of repeated disaccharides units of GlcA or IdoA and N-acetyl-galactosamine (GalNAc). The expression of both CS synthase CHSY1/CSS-1 and CHPF/CSS-2, which catalyze the polymerization of CS/DS, did not differ between CTR and DMD biopsies (Figs. 4A, B); this is consistent with the result in Figure 3C, which indicates that the length of the CS/DS chains was not modified in DMD. In addition, the levels of enzymes involved in GlcA/ IdoA modifications, UST (uronyl-2-sulfotransferase) and DSE (dermatan sulfate epimerase) were not different between CTR and DMD (Figs. 4C, D). Among the 3 different isoforms of the C4ST enzyme (CHST11/C4ST-1, CHST12/C4ST-2, and CHST13/C4ST-3), which catalyze the 4-O-sulfation of GalNAc (Figs. 4EYG), only CHST11 was significantly greater in DMD versus CTR biopsies (* p G 0.05) (Fig. 4E). This is in accordance with the increased amount of MonoS disaccharides observed in the DMD muscle biopsies (Fig. 3C). The expression of CHST14/D4ST-1 enzyme (which is responsible for the 4-O-sulfation of GalNAc adjacent to IdoA), CHST3/C6ST-1 and CHST7/C6ST-2 enzymes (which catalyze the 6-O-sulfation of GalNAc), and GalNAc4S-6ST/CHST15 enzyme (which transfers a sulfate group on GalNAc-4S) did not differ between DMD and CTR biopsies (Figs. 4HYK). Interestingly, the expression of the CS hydrolase HYAL-4/ CSHY enzyme, which is involved in the depolymerization of CS, was downregulated in DMD samples (** p G 0.01), suggesting diminished degradation of CS/DS in the skeletal muscle of DMD patients (Fig. 4L). All of these results are consistent with an increased level of sulfation caused by an increased proportion of MonoS disaccharides and a decreased turnover of CS/DS.
CS/DS Structural Modifications Because increased amounts of CS/DS, as measured by the 1Y9 dimethyl-methylene blue assay, may be attributable to an increase in GAG chain length and/or an increase in disaccharide unit sulfation, we assessed GAG chain length using agarose gel electrophoresis. We observed no difference in GAG chain lengths extracted from CTR and DMD biopsies (Fig. 3B). High-performance liquid chromatography analysis, however, allows one to define the relative proportion of each of the disaccharide units in the composition of CS/DS chains. In DMD patient biopsies, the proportion of nonsulfated disaccharides was significantly less than in CTR biopsies (36.0%
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DISCUSSION Extracellular matrix remodeling and fibrosis represent key components of the pathologic evolution in DMD. Although innovative therapeutic strategies are being tested to restore dystrophin expression (such as exon-skipping, gene therapy, or cell therapy), none of these will target the fibrosis that may limit access of the therapeutic agent to the muscle fibers. Fibrosis is a complex but not yet fully understood process characterized by excessive accumulation of collagens and ECM components (6). Recently, it was demonstrated in DMD that endomysial fibrosis was the only myopathologic parameter Ó 2014 American Association of Neuropathologists, Inc.
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J Neuropathol Exp Neurol Volume 73, Number 8, August 2014
Chondroitin Sulfate Remodeling in DMD
FIGURE 4. Analysis of enzymes involved in CS/DS metabolism. Analysis of enzymes involved in CS/DS metabolism in control (CTR) and DMD patient skeletal muscle biopsies by real-time RT-PCR (AYF, HYK) and RT-PCR (G, L). Values were normalized using B2M mRNA as reference. Each value is presented as mean T SEM. Student t-test was used to compare differences between CTR and DMD biopsies. Differences were considered significant at * p G 0.05 or ** p G 0.01. CHPF, chondroitin polymerizing factor; CHSY1, CS synthase-1; DSE, DS epimerase; GlcA, glucuronic acid; IdoA, iduronic acid; UST, uronyl-2-sulfotransferase.
that significantly correlated with poor motor outcome (30). Studying matrix components could help to understand mechanisms involved in the deposition of fibrosis and to find therapeutic targets to reverse fibrosis.
CS/DS, But Not HS, Are Increased in DMD This is the first study to investigate GAG content in DMD. The most striking and original finding in our study is the marked and selective increase in CS/DS in the muscles of DMD patients, whereas no changes in the amount of HS were observed between CTR and DMD biopsies; these results were found by both histochemical and biochemical quantification of GAGs. Advanced stages of DMD are characterized by an absence of muscle regeneration caused by a depletion of the satellite cell compartment in chronically damaged DMD muscle. To date, many studies have been conducted on syndecans, a family of PGs that carry both HS and CS chains, highlighting their roles as regulators of skeletal muscle regeneration, probably through modulation of signaling pathways by HS chains (17, 31, 32). It is possible that at this stage of the disease, HSmediated mechanisms, which participate in the regulation of muscle regeneration, are no longer active. The histochemical
approach localized CS both in the body of the ECM and in the basal lamina, whereas DS was in the endomysial space of the skeletal muscle. These results are in agreement with previous histochemical studies on PGs, which demonstrated that CS/DSPG biglycan and DS-PG decorin accumulate throughout the perimysium and endomysium (33). Recently, a study using a similar immunostaining approach performed on human skeletal muscle showed a decrease in HS and an increase in CS in the diaphragm muscles of patients affected by chronic obstructive pulmonary disease, which is associated with dysfunction of inspiratory muscles (34). Altogether, these results indicate that a selective increase in CS/DS content represents a specific response to a controlled mechanism that induces ECM remodeling in dystrophic and other pathologically altered muscles.
DMD Is Characterized by an Increased Proportion of CS/DS 4-O-Sulfation In addition to the accumulation of CS/DS in DMD, we demonstrate for the first time that GAG fine structure is also modified as well. We observed an enriched proportion of MonoS units and probably a specific increase in 4-O-sulfation of GalNAc residues, consistent with the upregulation of
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the CHST11/C4ST-1 enzyme. Our results are in agreement with a previous study, which suggested that the CHST11/ C4ST-1 isoform acts in cooperation with the chondroitin Nacetylgalactosaminyltransferase 2 to increase the amount of CS biosynthesized by cells (35). We also found that the expression of the CS hydrolase HYAL-4/CSHY enzyme, which is involved in the depolymerization of CS (36Y38), was downregulated in DMD samples. Because we found no modifications in GAG chain length in DMD, the increase in CS/DS in DMD biopsies can also be explained by an increase in the number of GAG chains, consistent with a diminished activity of HYAL-4/CSHY. Together, these results suggest that, in DMD patients, CS chains become increasingly sulfated and their turnover is reduced. Interestingly, analysis of CS/DS metabolic enzymes in the affected cricopharyngeal muscle of oculopharyngeal muscular dystrophy patients, where atrophy and fibrosis are present (39), showed a different regulation of the machinery of sulfation of CS/DS (data not shown). This indicates that the modifications in GAG sulfation observed in DMD are not a general muscle fibrotic process and could be specific for DMD. Recently, Mikami et al (40) showed that levels of CS chains in the murine myoblast cell line C2C12 are dramatically diminished at the stage of extensive syncytial myotube formation, indicating that a decrease in CS content, mediated by Hyal-1 activity, is required for muscle differentiation. In addition, they showed that enzymatic removal of all CS species improved the dystrophic pathology in the muscles of mdx mice, a model for DMD. Therefore, it would be of interest to determine if the upregulation of CS/DS and, more specifically, the increasing proportion of 4-O-sulfation in human dystrophic muscle are associated with changes in GAG functions. We have previously demonstrated that GAGs from an aged myocardium present altered sulfation patterns, leading to disruption of growth factor binding and activities (24). We showed that GAGs from aged cardiac muscle display better affinity for vascular endothelial growth factor, whereas aging significantly reduces the capacity of these GAGs to bind fibroblast growth factor-1 and fibroblast growth factor-2. Interestingly, TGFA (one of the key players in muscle tissue remodeling) (6), CTGF (41), and OPN (42) are all upregulated in DMD (41Y43), and each of them is a HBPs, which is regulated by GAG chains. In addition, growth factors of the TGFA superfamily have been identified as positive regulators of the expression of the CHST11/C4ST-1 gene (44), suggesting a possible positive feedback loop on fibrosis. In conclusion, we describe for the first time an accumulation of CS/DS GAGs in DMD muscle and a modification of the sulfation pattern of CS/DS in DMD biopsies, suggesting a possible role for 2 enzymes (CHST11/C4ST-1 and HYAL-4/CSHY) implicated in the machinery of sulfation of CS/DS. Understanding of the roles of GAGs in muscular dystrophies has only started to emerge. Thus, it remains to be determined if the alteration of GAGs is a consequence of the primary remodeling that contributes to worsening of the disease and then participates in the formation of fibrosis or, in contrast, if their modulation is a compensatory process. In DMD, further studies using new glycomic technologies are required to more finely characterize CS/DS functionality in
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muscle biopsies to determine the relationship between the changes occurring in CS/DS sulfation pattern and their ability to control such HBP-induced signaling, leading to the formation and maintenance of fibrosis. Finally, GAG fine structure characterization in DMD biopsies may provide a new tool for the design of a potential new class of anti-fibrotic drug candidates. ACKNOWLEDGMENTS We gratefully thank C. Trollet, A. Bigot, and P. Klein for fruitful discussions and critical reading of the manuscript. We also thank G. Butler-Browne and V. Mouly for their support and for precious discussions during the preparation of the manuscript. We thank S. Vasseur and M. Chapart from Myobank for providing muscle biopsies. We thank OTR3 (Paris, France) for assuring M. Gilbert’s salary independently of this work. REFERENCES 1. Mendell JR, Shilling C, Leslie ND, et al. Evidence-based path to newborn screening for Duchenne muscular dystrophy. Ann Neurol 2012;71:304Y13 2. Hoffman EP, Brown RH Jr, Kunkel LM. Dystrophin: The protein product of the Duchenne muscular dystrophy locus. Cell 1987;51:919Y28 3. Emery AE. The muscular dystrophies. Lancet 2002;359:687Y95 4. Blake DJ, Weir A, Newey SE, et al. Function and genetics of dystrophin and dystrophin-related proteins in muscle. Physiol Rev 2002;82:291Y329 5. Durbeej M, Campbell KP. Muscular dystrophies involving the dystrophinglycoprotein complex: An overview of current mouse models. Curr Opin Genet Dev 2002;12:349Y61 6. Serrano AL, Munoz-Canoves P. Regulation and dysregulation of fibrosis in skeletal muscle. Exp Cell Res 2010;316:3050Y58 7. Gargioli C, Coletta M, De Grandis F, et al. PlGF-MMP-9Yexpressing cells restore microcirculation and efficacy of cell therapy in aged dystrophic muscle. Nat Med 2008;14:973Y78 8. Le Hir M, Goyenvalle A, Peccate C, et al. AAV genome loss from dystrophic mouse muscles during AAV-U7 snRNA-mediated exon-skipping therapy. Mol Ther 2013;21:1551Y58 9. Raghow R. The role of extracellular matrix in postinflammatory wound healing and fibrosis. FASEB J 1994;8:823Y31 10. Frantz C, Stewart KM, Weaver VM. The extracellular matrix at a glance. J Cell Sci 2010;123:4195Y200 11. Sugahara K, Mikami T, Uyama T, et al. Recent advances in the structural biology of chondroitin sulfate and dermatan sulfate. Curr Opin Struct Biol 2003;13:612Y20 12. Volpi N. Therapeutic applications of glycosaminoglycans. Curr Med Chem 2006;13:1799Y810 13. Jenniskens GJ, Veerkamp JH, van Kuppevelt TH. Heparan sulfates in skeletal muscle development and physiology. J Cell Physiol 2006;206: 283Y94 14. Zhang L. Glycosaminoglycan (GAG) biosynthesis and GAG-binding proteins. Prog Mol Biol Transl Sci 2010;93:1Y17 15. Handel TM, Johnson Z, Crown SE, et al. Regulation of protein function by glycosaminoglycansVas exemplified by chemokines. Annu Rev Biochem 2005;74:385Y410 16. Friedl A, Chang Z, Tierney A, et al. Differential binding of fibroblast growth factor-2 and -7 to basement membrane heparan sulfate: Comparison of normal and abnormal human tissues. Am J Pathol 1997;150: 1443Y55 17. Pisconti A, Bernet JD, Olwin BB. Syndecans in skeletal muscle development, regeneration and homeostasis. Muscles Ligaments Tendons J 2012;2:1Y9 18. Caceres S, Cuellar C, Casar JC, et al. Synthesis of proteoglycans is augmented in dystrophic mdx mouse skeletal muscle. Eur J Cell Biol 2000;79:173Y81 19. Casar JC, McKechnie BA, Fallon JR, et al. Transient up-regulation of biglycan during skeletal muscle regeneration: Delayed fiber growth along with decorin increase in biglycan-deficient mice. Dev Biol 2004;268: 358Y71
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