Antxr2-mediated fine-tuning of Collagen VI ensures skeletal muscle function

Antxr2-mediated fine-tuning of Collagen VI ensures skeletal muscle function | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Antxr2-mediated fine-tuning of Collagen VI ensures skeletal muscle function Samuele Metti , View ORCID Profile Lucie Josepha Bracq , Matteo Signor , Nicola Facchinello , Lisa Gambarotto , View ORCID Profile Leonardo Nogara , Giulia Pigato , Laurence Abrami , Béatrice Kunz , Julien Duc , Patrizia Sabatelli , Aurora De Acutis , Gabriele Maria Fortunato , Giovanni Vozzi , View ORCID Profile Paola Braghetta , View ORCID Profile Bert Blaauw , View ORCID Profile F. Gisou van der Goot , View ORCID Profile Paolo Bonaldo doi: https://doi.org/10.1101/2025.09.11.675515 Samuele Metti a Department of Molecular Medicine, University of Padova , Padova, Italy b Global Health Institute, School of Life Sciences, Ecole Polytechnique Fédérale de Lausanne , Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: bonaldo{at}bio.unipd.it Lucie Josepha Bracq b Global Health Institute, School of Life Sciences, Ecole Polytechnique Fédérale de Lausanne , Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Lucie Josepha Bracq Matteo Signor a Department of Molecular Medicine, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Nicola Facchinello c Neuroscience Institute, Italian Research Council (CNR) , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Lisa Gambarotto a Department of Molecular Medicine, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Leonardo Nogara d Department of Biomedical Sciences, University of Padova , Padova, Italy e Department of Pharmaceutical and Pharmacological Sciences, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Leonardo Nogara Giulia Pigato a Department of Molecular Medicine, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Laurence Abrami b Global Health Institute, School of Life Sciences, Ecole Polytechnique Fédérale de Lausanne , Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site Béatrice Kunz b Global Health Institute, School of Life Sciences, Ecole Polytechnique Fédérale de Lausanne , Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site Julien Duc b Global Health Institute, School of Life Sciences, Ecole Polytechnique Fédérale de Lausanne , Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site Patrizia Sabatelli f CNR, Institute of Molecular Genetics “Luigi Luca Cavalli-Sforza”, Unit of Bologna , Bologna, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Aurora De Acutis g Department of Information Engineering and Research Centre “E. Piaggio”, University of Pisa , Pisa, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Gabriele Maria Fortunato g Department of Information Engineering and Research Centre “E. Piaggio”, University of Pisa , Pisa, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Giovanni Vozzi g Department of Information Engineering and Research Centre “E. Piaggio”, University of Pisa , Pisa, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site Paola Braghetta a Department of Molecular Medicine, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Paola Braghetta Bert Blaauw d Department of Biomedical Sciences, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Bert Blaauw F. Gisou van der Goot b Global Health Institute, School of Life Sciences, Ecole Polytechnique Fédérale de Lausanne , Lausanne, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for F. Gisou van der Goot For correspondence: bonaldo{at}bio.unipd.it Paolo Bonaldo a Department of Molecular Medicine, University of Padova , Padova, Italy Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Paolo Bonaldo For correspondence: bonaldo{at}bio.unipd.it Abstract Full Text Info/History Metrics Preview PDF ABSTRACT Tissue function relies on the extracellular matrix (ECM) that surrounds cells, providing structural and biochemical support. The complex ECM composition depends on an adequately tuned balance between the deposition and degradation of each of its components. Disequilibrium may cause disease, as observed for Collagen VI (COL6), for which mutations lead to muscular dystrophy. Here, we investigated the role of Anthrax Toxin Receptor 2 (ANTXR2/CMG2), a receptor that controls the turnover of COL6, in skeletal muscle. We show that ANTXR2 is mostly expressed by fibro-adipogenic precursors and that its deficiency in ANTXR2 null ( Antxr2 -/- ) mice leads to a premature and irregular COL6 accumulation in intramuscular connective tissue. This results in tissue stiffening and gradual, non-functional muscle hypertrophy, marked by impaired locomotion and myopathic signs. Our findings further indicate that COL6 accretion drives these alterations, as revealed by Antxr2 -/- ::Col6a1 -/- double knockout mice, highlighting the essential role of ANTXR2-mediated COL6 remodeling in maintaining ECM homeostasis and muscle functionality. SIGNIFICANCE STATEMENT Remodelling of the extracellular matrix (ECM) was long thought to rely almost exclusively on extracellular proteases. Increasing evidence, however, indicates that some ECM components may undergo intracellular degradation following receptor-mediated endocytosis, as we have found for Collagen VI (COL6). Here, we identify the COL6 receptor ANTXR2 as a critical regulator of ECM turnover in skeletal muscle.. When ANTXR2 is absent, COL6 builds up, followed by the accumulation of fibrillar collagens, without changes in gene expression. These alterations in muscle ECM lead to increased stiffness, myofiber defects and impaired locomotor activity. Our findings establish ANTXR2 as a key regulator in ECM remodelling, offering new insights into potential treatments for conditions associated with defective ECM remodeling, such as aging and congenital muscular dystrophies. INTRODUCTION The extracellular matrix (ECM), an intricate meshwork of proteins and proteoglycans, is required to organize cells into tissues and maintain organ structure. Such macromolecular network regulates a number of cellular functions and processes through biochemical and biomechanical cues 1 , 2 . The ECM is highly dynamic, being remodelled according to specific needs through a fine-tuned balance of protein synthesis and assembly, post-translational modifications, recycling and degradation. Failure of this tight regulation contributes to a range of disease conditions such as cancer progression, osteoarthritis, and muscular dystrophies 3 – 5 . The mechanisms underlying the control of ECM remodelling in the different tissues are however incompletely understood. ECM degradation can occur through two main routes 5 , 6 . The first and most documented is extracellular degradation, which is mainly orchestrated by different families of secreted proteases whose activity is responsible for the cleavage of ECM components within the extracellular space 7 , 8 . The second mode of degradation is intracellular, following receptor-mediated endocytosis of ECM macromolecules and lysosomal clearance 9 , 10 . Intracellular degradation has been largely overlooked due to the limited characterization of the involved molecular players. In this study, we have investigated the relevance of receptor-mediated clearance of Collagen VI (COL6) in skeletal muscle, given the key role of this ECM component for proper muscle function. In past work, we identified Anthrax Toxin Receptor 2 (ANTXR2), also known as Capillary Morphogenesis Gene 2 (CMG2), as a cell surface receptor involved in the homeostasis of COL6, by mediating COL6 endocytosis and lysosomal delivery 11 , 12 . The ANTXR2 name stems from the well-known role of this surface protein in binding and internalizing Bacillus anthracis toxins 13 . The physiological roles of ANTXR2 are only beginning to be elucidated 14 . Loss-of-function mutations in the human ANTXR2 gene cause hyaline fibromatosis syndrome (HFS, MIM #228600) 15 , a severe and debilitating autosomal recessive disease characterized by the formation of non-cellularized subcutaneous and visceral hyalinized nodules highly enriched in COL6 11 . This human condition is mirrored to some extent by ANTXR2 null ( Antxr2 -/- ) mice, which progressively accumulate COL6 in the uterus, affecting organ contraction, thus leading to the inability to deliver pups 11 , 16 . COL6 is made of three major polypeptide chains [α1(VI), α2(VI), α3(VI)] coded by separate genes ( COL6A1 , COL6A2 and COL6A3 in humans) 17 . It is unique within the collagen superfamily because it forms a distinctive network of beaded microfilaments in the ECM of various organs, where it binds several ECM components, such as fibrillar collagens 18 , fibronectin 19 , 20 and proteoglycans 21 . The proper deposition of COL6 in the ECM is particularly critical for skeletal muscle, and COL6 deficiency due to mutations in its genes is causative for inherited muscle disorders known as COL6-related myopathies 22 – 24 . COL6 synthesis and matrix deposition are finely regulated in different tissues 25 – 27 , however the mechanisms that control its turnover in muscle are largely unknown. In this study, we uncover a critical role of ANTXR2 in regulating COL6 turnover in skeletal muscle by exploiting an ANTXR2-deficient mouse model. We find that ANTXR2 is predominantly expressed by fibro-adipogenic precursors (FAPs) and that its genetic ablation leads to an early onset and progressive buildup of COL6 in muscle ECM, with a subsequent increase in fibrillar collagens accompanied by myofiber changes. Notably, although Antxr2 -/- muscles display hypertrophy, the added muscle mass is non-functional and exhibits hallmarks of myopathy. To determine causality, we generated and performed phenotypic characterization of Antxr2 -/- ::Col6a1 -/- double knockout mice, which provided a genetic proof that excessive COL6 is the key driver of the pathological features observed in Antxr2 -/- skeletal muscle. This work demonstrates that the proper regulation of COL6 homeostasis by ANTXR2 is critical for preserving ECM features and, consequently, muscle structure and function. RESULTS Antxr2 is expressed by FAPs in adult skeletal muscle We first evaluated the expression of Antxr2 gene in the different cell populations of adult mouse muscles. We mined the GSE147127 dataset containing single-nucleus RNA-seq data of skeletal muscles from C57BL/6 mice of different ages 28 . Analysis of tibialis anterior (TA) muscles from 5-month-old mice revealed that Antxr2 expression is high in FAPs, whereas it is extremely low in the nuclei of myofibers, also know as myonuclei ( Fig. 1A ). Expression in FAPs and very low expression in myofibers was also observed for the three major COL6 genes, Col6a1 , Col6a2 and Col6a3 genes ( Fig. 1A ), consistent with previous findings that they are predominantly expressed by muscle fibroblasts, rather than by mature myofibers 27 – 29 . Download figure Open in new tab Figure 1. ANTXR2 is mainly expressed by FAPs in adult skeletal muscle. ( A ) Dot plot showing the expression of Antxr2 , Col6a1 , Col6a2 and Col6a3 genes in the different cell populations of TA muscles from 5-month-old C57BL/6 WT mice, based on available single-nucleus RNA-seq data deposited in the GSE147127 dataset. Dot size reflects the percentage of cells with a non-zero expression level and color scale represents the average expression level across all cells within the cluster. MTJ, myotendineous junction; MuSCs, muscle stem cells; NMJ, neuromuscular junction. (B) Representative fluorescence microscopy images of in situ hybridization for Antxr2 mRNA in TA cross-sections of 8-month-old WT animals, showing colocalization of Antxr2 mRNA (magenta) with the FAP marker Vimentin (VIM, green). Nuclei were stained with Hoechst (blue). Muscle fibers are outlined with white dashed lines. White arrowheads point at Vimentin-positive cells. The violin plot shows the number of Antxr2 mRNA positive dots localized in close proximity of Vimentin positive (VIM+) or negative (VIM–) regions ( n = 20 fields from 3 WT mice; **, P <0.01; two-tailed Mann-Whitney test). Scale bar, 10 μm. ( C ) Representative confocal images of immunostaining for Vimentin (VIM, green), ANTXR2 (magenta), and COL6 α3 (α3(VI), yellow) in EDL myofiber bundles of 8-month-old WT and Antxr2 -/- animals. Muscle fibers are outlined with white dashed lines. Scale bar, 10 μm. To corroborate the RNA-seq data, we performed RNA in situ hybridization in TA cross-sections from 8-month-old wild-type (WT) animals. Antxr2 mRNA was indeed predominantly found in FAPs, positive for the fibroblast marker Vimentin ( Fig. 1B ). Immunofluorescence of myofiber bundles isolated from Extensor Digitorum Longus (EDL) muscles confirmed ANTXR2 staining in Vimentin-positive FAPs in WT samples, whereas no ANTXR2 signal was detectable in Antxr2 -/- samples ( Fig. 1C ). Staining with an antibody against the α3(VI) polypeptide showed COL6 localization in the external layer of the basement membrane ( Fig. 1C ). No signal for ANTXR2 protein was found in WT myofibers ( Fig. 1C ), nor were any Antxr2 mRNA-positive dot detected in the proximity of myonuclei (Supplementary Fig. 1). Together, these data show that FAPs are the major cell population expressing ANTXR2 within adult skeletal muscle. Abnormal progressive buildup of COL6 in ANTXR2-deficient muscles Given the ability of ANTXR2 to bind COL6 and mediate its endocytosis and degradation in lysosomes in a primary fibroblast culture system 11 , 12 , we assessed COL6 abundance in muscles of WT and Antxr2 -/- mice at different postnatal ages. Western blotting for the COL6 α1(VI) and α3(VI) chains in protein lysates of gastrocnemius muscle from 5-day-old pups did not reveal any significant differences between WT and Antxr2 -/- samples ( Fig. 2A ). To corroborate these findings, we performed immunofluorescence for the COL6 α3(VI) and α6(VI) chains, taking into account previous studies which showed that the α3(VI) polypeptide can be substituted by other minor COL6 chains in some conditions and tissues, and in particular α6(VI) in skeletal muscles 30 , 31 . For both chains, immunostaining was similar between WT and Antxr2 -/- muscles, with COL6 α3(VI) restricted to perimysium (the thicker connective tissue layer encasing muscle fiber bundles), whereas COL6 α6(VI) was also abundant in the endomysial layer surrounding individual myofibers ( Fig. 2B ). These biochemical and immunohistochemical data indicate that ANTXR2 ablation does not visibly affect COL6 abundance and localization in skeletal muscle during early postnatal life. Download figure Open in new tab Figure 2. COL6 undergoes progressive accretion in ANTXR2-deficient muscles. ( A ) Western blotting for COL6 α1 [α1(VI)] and α3 [α3(VI)] chains in protein extracts from gastrocnemius muscles of 5-day-old (P5) WT and Antxr2 −/− mice. Vinculin was used as a loading control. The histograms on the right show the densitometric quantification of α1(VI) and α3 (VI) signals, normalized on Vinculin protein levels. Data are expressed as mean ± s.e.m. ( n = 5-6, each group; NS, not significant; two-tailed Mann-Whitney test). ( B, D-F ) Representative confocal images of immunostaining for COL6 α3 (α3(VI), magenta) and α6 (α6(VI), green) chains in cross-sections of gastrocnemius muscles from 5-day-old WT and Antxr2 −/− mice ( B ), and in cross-sections of TA muscle from 1-month-old ( D ), 3-month-old ( E ), and 8-month-old ( F ) WT and Antxr2 −/− mice. Note that, as expected, myofiber size increases after birth and along early postnatal life, being larger in 3-and 8-month-old when compared to 5-day-old muscle sections. The respective histograms on the right show the percentage of threshold area positive for α3(VI) or α6(VI) staining. Scale bars, 50 μm. Data are expressed as mean ± s.e.m. ( n = 4–8, each group; NS, not significant; *, P <0.05; **, P <0.01; ***, P <0.001; two-tailed Mann-Whitney test). (C) Western blotting for COL6 α1 [α1(VI)] and α3 [α3(VI)] chains in protein extracts from TA muscles of 1-, 3-and 8-month-old (mo) WT and Antxr2 −/− mice. Vinculin was used as a loading control. The histograms on the bottom show the densitometric quantification of α1(VI) (left panel) and α3(VI) (right panel) signals, normalized on Vinculin protein levels. Data are expressed as mean±s.e.m. ( n = 4–6, each group; *, P <0.05; ***, P <0.001; ****, P <0.0001; two-way ANOVA test, with Sidak’s test for multiple comparisons). ( G ) RT-qPCR analysis for Col6a1 , Col6a2 and Col6a3 transcript levels in gastrocnemius muscles of 8-month-old WT and Antxr2 −/− mice. Gapdh was used as housekeeping gene. Data are expressed as mean ± s.e.m. ( n = 9–10, each group; NS, not significant; two-tailed Mann-Whitney test). a.u., arbitrary units; VNC, vinculin; GA, gastrocnemius. Conversely, at 1 month of age Antxr2 -/- mice displayed a progressive accumulation of COL6 chains, as revealed by western blotting of TA muscles, becoming increasingly more abundant in 3-and 8-month-old mice when compared to the corresponding samples from age-matched WT animals ( Fig. 2C ). Immunofluorescence of TA muscles at the same postnatal ages confirmed markedly increased of both α3(VI) and α6(VI) in the endomysium of Antxr2 -/- animals ( Fig. 2D -F). A similar pattern of COL6 accumulation was displayed by the Gastrocnemius of 8-month-old Antxr2 -/- mice (Supplementary Fig. 2A, B). To assess whether COL6 protein accumulation was a consequence of increased gene expression, we quantified transcript levels. RT-qPCR analyses in TA of 8-month-old animals showed no significant difference in Col6a1 , Col6a2 and Col6a3 mRNA levels between WT and Antxr2 -/- mice ( Fig. 2G ). We next considered whether COL6 assembly and secretion could be affected. Past studies demonstrated that COL6 undergoes a distinctive stepwise intracellular assembly process, in which three chains associate into monomers (∼500 kDa), dimers (∼1,000 kDa) and tetramers (∼2,000 kDa), which are finally secreted and deposited in the ECM 17 . Protein electrophoresis into composite acrylamide-agarose gels under non-reducing conditions showed that all COL6 species increased proportionally in TA lysates of Antxr2 -/- mice, indicating that the assembly of monomers, dimers and tetramers is not appreciably affected (Supplementary Fig. 2C). Interestingly, the high COL6 content of Antxr2 -/- muscles led to a significant increase in the SDS-insoluble COL6 fraction (Supplementary Fig. 2D), suggestive of increased crosslinking. Altogether, these data show that ANTXR2 null muscles undergo a progressive buildup of COL6 in the ECM, despite normal expression of COL6 genes and proper COL6 biosynthesis with the expected formation of monomers, dimers and tetramers. The greatly increased abundance of COL6 and its increased SDS-insolubility in Antxr2 -/- samples therefore support a critical role for ANTXR2 in regulating COL6 turnover in skeletal muscle. Fibrillar collagens accumulate in muscles of adult ANTXR2-deficient mice We analyzed the tissue architecture of TA muscles from 8-month-old Antxr2 -/- mice and found that they did not overtly differ from that of WT mice ( Fig. 3A ). In particular, histological analysis by haematoxylin-eosin staining did not reveal any sign of fibrous-adipose replacement or cellular infiltration, which would be expected in the presence of active fibrosis or necrotic lesions 32 . However, Antxr2 -/- muscles repeatedly displayed the presence of cytoplasmic inclusions within myofibers, which were not present in the corresponding WT samples ( Fig. 3A , white arrowheads). The histological profile of such abnormalities is reminiscent of tubular aggregates, one of the most recurrent myofiber alterations found in neuromuscular diseases 33 , 34 . This was confirmed by Gomori trichrome staining, which revealed a markedly increased incidence of myofibers containing staining-positive aggregates in 8-month-old Antxr2 -/- male mice when compared to age-and sex-matched WT samples ( Fig. 3B, C ), predominantly affecting larger glycolytic myofibers. Electron microscopy revealed that these cytoplasmic inclusions were membranous accumulations characterized by tubular and saccular dilations, thereby definitely identifying them as tubular aggregates ( Fig. 3D ). These stained positively for the sarco/endoplasmic reticulum markers Calsequestrin, STIM1 and SERCA1/2/3, and the T-tubule markers Orai1 and DHPR (Supplementary Fig. 3A-C). Altogether, this analysis shows that TA muscles from 8-month-old Antxr2 -/- mice overal maintain a proper architecture, but display signs of alterations as exemplified by the presence of tubular aggregates. Download figure Open in new tab Figure 3. Muscles from adult ANTXR2-deficient mice display changes in myofibers and in the ECM. ( A ) Representative images of cross-sections of fresh frozen TA muscles from 8-month-old WT and Antxr2 −/− mice following staining with hematoxylin-eosin. White arrowheads indicates cytoplasmic inclusions. Scale bar, 50 μm. ( B ) Modified Gomori trichrome staining of cross-sections of fresh frozen TA muscles from 8-month-old WT and Antxr2 -/- male mice. Gomori-positive cytoplasmic inclusions (red), corresponding to tubular aggregates, are present in Antxr2 -/- muscles but not in WT muscles. The panels on the right show higher magnification images of Antxr2 -/- muscles, displaying different morphologies of tubular aggregates (intermyofibrillar, subsarcolemmal and fragmented). White asterisks indicate smaller oxidative fibers. Scale bar, 50 μm. ( C ) Quantification of the percentage of vacuolated myofibers, determined by modified Gomori trichrome staining as in (B). Data are expressed as mean ± s.e.m. ( n = 4-5, each group; *, P <0.05; two-tailed Mann-Whitney test). ( D ) Representative transmission electron microscopy images of ultrathin sections of TA muscles from 8-month-old Antxr2 −/− male mice. Areas displaying tubular aggregates are highlighted in light blue. Scale bar, 1 μm. ( E ) Representative images of cross-sections of fresh frozen TA muscles from 8-month-old WT and Antxr2 −/− mice following staining with Sirius Red. Scale bar, 50 μm. ( F ) Top panel, histograms showing the percentage of threshold area positive for Sirius Red (SR, acquired under polarized light). Data are expressed as mean ± s.e.m. ( n = 5, each group; **, P <0.01; two-tailed Mann-Whitney test). Bottom panel, Tukey box-and-whisker plots showing the total hydroxyproline (Hyp) content of gastrocnemius muscles from 8-month-old WT and Antxr2 −/− mice ( n = 10, each group; *, P <0.05; two-tailed Mann-Whitney test). ( G ) Representative transmission electron microscopy images of TA ultrathin sections from 8-month-old WT and Antxr2 −/− mice. The endomysial area is highlighted in light blue. An abnormal accumulation of fibrillar collagens (orange arrowheads) is clearly visible in Antxr2 −/− muscles. Scale bar, 500 nm. ( H ) Representative confocal images of COL1 (white, left panels) and COL3 (white, right panels) immunostaining in cross-sections of TA muscles from 8-month-old WT and Antxr2 −/− mice. The histograms on the right show the percentage of threshold area positive for COL1 (top panel) and COL3 (bottom panel) staining. Data are expressed as mean ± s.e.m. ( n = 4–5, each group; *, P <0.05; **, P <0.01; two-tailed Mann-Whitney test). Scale bar, 50 μm. ( I ) Western blotting (upper panel) and relative densitometric quantification (lower panel) of COL3 content in protein extracts from TA muscles of 8-month-old WT and Antxr2 −/− mice. Vinculin (VNC) was used as a loading control. Data are expressed as mean ± s.e.m. ( n = 4–5, each group; **, P <0.01; two-tailed Mann-Whitney test). ( J ) Histogram showing the quantification of the elastic modulus of fresh TA muscles from 8-month-old WT and Antxr2 −/− mice. Data are expressed as mean ± s.e.m. ( n = 6, each group; **, P <0.01; two-tailed Mann-Whitney test). Since biochemical and immunofluorescent analysis of Antxr2 -/- skeletal muscle revealed a progressive buildup of COL6, we next investigated whether this was accompanied by alterations in other ECM components. Sirius red staining showed increased deposition of fibrillar collagens in the endomysium of TA muscles from 8-month-old Antxr2 -/- mice when compared to age-matched WT mice ( Fig. 3E, F ), a difference that was not observed in younger mice (Supplementary Fig. 4A, B). The increase in fibrillar collagens at 8 months, but not earlier, in TA muscles from Antxr2 -/- mice was confirmed biochemically using hydroxyproline quantification, an amino acid signature for these ECM structures ( Fig. 3F and Supplementary Fig. 4B). Finally, the increased abundance of collagen fibrils in TA muscles was readily detectable by electron microscopy analysis of longitudinal ultrathin sections, in the vicinity of the myofiber basal lamina of ANTXR2-deficient muscles ( Fig. 3G ). We next specifically monitored Collagen I (COL1) and Collagen III (COL3), the two most abundant fibrillar collagens in skeletal muscle, using immunofluorescence and western blotting. Both COL1 and COL3 were increased in TA and Gastrocnemius muscles of 8-month-old Antxr2 -/- animals when compared to age-matched WT samples ( Fig. 3H, I and Supplementary Fig. 4C). An increased abundance of these fibrillar collagens in TA muscles of Antxr2 -/- mice was already apparent at 3 months of age (Supplementary Fig. 4D). These data indicate that the progressive extracellular accumulation of collagens in muscles lacking ANTXR2 extends to COL1 and COL3, albeit it seems to occur subsequently to COL6 accumulation during postnatal growth and maturity. Interestingly, the ECM changes elicited by ANTRX2 ablation do not appear to impinge on muscle basal lamina, i.e. the specialized innermost ECM of muscle endomysium in contact with the plasma membrane of myofibers. Indeed, immunofluorescence for Perlecan (HSPG2), Collagen IV (COL4) and Fibronectin (FN1), three major components of muscle basal lamina 35 , appeared similar in their amount and localization in muscles of 3-and 8-month-old WT and Antxr2 -/- mice (Supplementary Fig. 5A-C). To determine whether the accumulation of fibrillar collagens was associated with an increased expression of their genes, we quantified by RT-qPCR the mRNA levels for Col1a1 and Col3a1 . No significant differences in COL1 and COL3 expression were observed between 8-month-old WT and Antxr2 -/- TA (Supplementary Fig. 4E), nor between sorted FAPs isolated from 8-month-old WT and Antxr2 -/- muscles (Supplementary Fig. 4F). Moreover, TA and FAPs did no display any significant difference in the expression of the pro-fibrogenic Acta2 and Lox genes between WT Antxr2 -/- animals (Supplementary Fig. 4E, F). Therefore, the accumulation of ECM observed follows a mechanism that differs from classical fibrosis, where increases in the expression of collagen genes are observed 32 . A typical consequence of increased collagen deposition is tissue stiffening 36 , 37 . We therefore measured the micromechanical properties of fresh skeletal muscle through a uniaxial tensile test, and found that 3-and 8-month-old Antxr2 -/- TA muscles displayed a significantly higher elastic modulus when compared to the corresponding WT muscles, a clear sign of increased stiffness ( Fig. 3J and Supplementary Fig. 4G). Taken together, these data indicate that ANTXR2 deficiency triggers a progressive buildup of COL6 and fibrillar collagens in the interstitial ECM and increased muscle stiffness, without any evidence of increased expression of their genes, highlighting a different mechanism of ECM alteration than the typical fibrotic process. ANTXR2-deficient muscles develop progressive hypertrophy We next assessed whether the ECM alterations detected in ANTXR2-deficient muscles affect myofiber organization. Antxr2 -/- mice progressively developed heavier muscles than WT mice, a difference that became evident at 8 months of age, despite similar whole body weight ( Fig. 4A and Supplementary Fig. 6A). Morphometric analyses of TA cross-sections showed a lower myofiber density ( Fig. 4B, C ), a significantly increased myofiber mean cross-sectional area (CSA; Fig. 4D , left panel) and mean minimum Feret diameter (MFD; Supplementary Fig. 6B, left panel) than age-matched WT. The difference became even more apparent when comparing the distribution of myofiber size ( Fig. 4D and Supplementary Fig. 6B, right panels). Differences in CSA and MFD distribution were already apparent in 3-month-old mice, despite similar mean values (Supplementary Fig. 6C, D). Download figure Open in new tab Figure 4. Muscles from adult ANTXR2-deficient mice are hypertrophic. ( A ) Normalized weight of TA, gastrocnemius (GA) and quadriceps (QUA) muscles (left panel) and total body weight (right panel) of 8-month-old WT and Antxr2 −/− mice. The weight of right and left muscles was averaged and normalized to body weight. Data are expressed as mean ± s.e.m. ( n = 16–31, each group; NS, not significant, **, P <0.01; two-tailed Mann-Whitney test). ( B ) Representative confocal images of wheat germ agglutinin (WGA, green) and type 2A myofibers (positive for Myosin IIA, red) in cross-sections of TA muscles from 3-month-old wild-type and Antxr2 −/− mice. Scale bar, 50 μm. ( C ) Histograms showing the myofiber density (number of myofibers per area unit) in TA muscles of 8-month-old WT and Antxr2 −/− mice. Data are expressed as mean±s.e.m. ( n = 5–6, each group; **, P <0.01; two-tailed Mann-Whitney test). (D) Left panel, mean CSA of myofibers of all types (All), of myofibers positive for myosin IIA (2A) and of myofibers negative for myosin IIA (2X2B) in TA muscles of 8-month-old WT and Antxr2 −/− male mice. Right panel, distribution of myofibers with different CSA (displayed as the percentage of total myofibers) in TA muscles of 8-month-old WT and Antxr2 −/− mice. Data are expressed as mean ± s.e.m. ( n = 5–6, each group; NS, not significant; *, P <0.05; **, P <0.01; two-tailed Mann-Whitney test). ( E ) Histogram showing the percentage of type 2A myofibers in TA muscles of 8-month-old WT and Antxr2 −/− mice. Data are expressed as mean ± s.e.m. ( n = 5-7, each group; NS, not significant; two-tailed Mann-Whitney test). ( F ) Western blotting (left panel) and relative densitometric quantification (right panel) showing puromycine incorporation by SUnSET assay in TA muscles of 8-month-old WT and Antxr2 −/− mice. Vinculin was used as a loading control. Data are expressed as mean ± s.e.m. ( n = 5, each group; *, P <0.05; two-tailed Mann-Whitney test). a.u., arbitrary units; VNC, vinculin. The above data indicated that the absence of ANTXR2 leads to myofiber hypertrophy. Such hypertrophy often involves a switch from small oxidative myofibers (type 2A fibers, positive for myosin IIA) to large glycolytic myofibers (type 2X and 2B fibers, negative for myosin IIA) 38 . Immunostaining of TA muscles from 8-month-old mice for Myosin IIA, however, showed that the ratio of 2A to 2X/2B myofibers was not altered in Antxr2 -/- animals ( Fig. 4E ). The 2X/2B fibers in Antxr2 -/- TA muscles had significantly increased mean CSA and MFD ( Fig. 4D and Supplementary Fig. 6B). Thus, the progressively hypertrophic phenotype was triggered in the more glycolytic myofibers. Since myofiber hypertrophy is generally associated with an increase in protein synthesis, we measured protein synthesis in vivo using the puromycin-based SUnSET assay 39 . Animals at 8 months of age were injected with puromycin and, following muscle tissue isolation, puromycin-labelled truncated peptides were detected by western blotting. Puromycin incorporation into nascent polypeptides was significantly higher in TA muscles of 8-month-old Antxr2 -/- mice when compared to age-matched WT samples ( Fig. 4F ). Altogether, these data show that skeletal muscles of ANTXR2-deficient mice undergo a progressive hypertrophy of glycolytic myofibers, associated with an increase in protein synthesis. COL6 buildup in the ECM is the driver of muscle alterations in Antxr2 -/- mice In past studies, we showed that female ANTXR2-deficient mice suffer from severe accumulation of ECM in the uterus and found that this phenotype could be fully reverted by the additional knockout of the Col6a1 gene, restoring female fertility 11 . We tested whether ablation of COL6 expression would similarly revert the myofiber hypertrophy displayed by ANTXR2-deficient muscles. Quantifications of CSA and MFD in cross-sections of TA ( Fig. 5A, B ) and Quadriceps ( Fig. 5C ) muscles from 8-month-old Antxr2 -/- :: Col6a1 -/ - mice showed that the mean CSA and MFD were significantly lower than for WT mice, in fact very similar to what can be observed for Col6a1 -/- mice ( Fig. 5A-C ). This was apparent when analysing the distribution of fiber CSAs, particularly for the TA, with a clear shift to smaller cross-sectional areas for fibers from mice lacking COL6, irrespective of the presence of ANTXR2 ( Fig. 5B, C ). Thus, the progressive buildup of COL6 is pivotal for the myofiber hypertrophy displayed by Antxr2 -/- muscles. Download figure Open in new tab Figure 5. COL6 drives fibrillar collagen accumulation and myofiber hypertrophy in ANTXR2-deficient muscles. ( A ) Representative images of paraffin-embedded TA muscle cross-sections from 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- mice stained with hematoxylin-eosin (H&E). White arrowheads indicate small atrophic fibers. Scale bar, 50 μm. ( B ) Left panels, mean CSA and MFD of TA myofibers of 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- female mice. Right panel, distribution of myofibers with different CSA (displayed as the percentage of total myofibers) in TA muscles of 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- female mice. Data are expressed as mean ± s.e.m. ( n = 3, each group; NS, not significant; *, P <0.05; one way ANOVA, with Dunnet’s test for multiple comparison). ( C ) Left panels, mean CSA and MFD of quadriceps myofibers of 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- male mice. Right panel, distribution of myofibers with different CSA (displayed as the percentage of total myofibers) in quadriceps muscles of 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- male mice. Data are expressed as mean ± s.e.m. ( n = 3, each group; NS, not significant; *, P <0.05; ** P <0.01; one way ANOVA, with Dunnet’s test for multiple comparison). ( D ) Left panel, representative images of paraffin-embedded TA muscle cross-sections from 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- mice stained with Sirius Red (SR). Right panel, Histograms showing the percentage of threshold area positive for Sirius Red in cross-sections of TA (left histogram) and quadriceps (right histogram) muscles from 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- mice. Data are expressed as mean ± s.e.m. ( n = 3, each group; NS, not significant; *, P <0.05; **, P <0.01; ***, P <0.001; one way ANOVA, with Dunnet’s test for multiple comparison). Scale bar, 50 μm. ( E ) Representative confocal images of COL3 (white, left panels) immunostaining in TA cross-sections from 8-month-old WT, Col6a1 -/- and Antxr2 -/- :: Col6a1 -/- mice. The histograms on the right show the percentage of threshold area positive for COL3 staining. Data are expressed as mean ± s.e.m. ( n = 3, each group; NS, not significant; ***, P <0.001; one-way ANOVA, with Dunnet’s test for multiple comparison). Scale bar, 50 μm. We also tested whether the excessive deposition of fibrillar collagens observed in Antxr2 -/- muscles would be reverted by the absence of COL6. Sirius red staining of TA and Quadriceps ( Fig. 5D ) and COL3 immunostaining of TA ( Fig. 5E ) of 8-month-old Antxr2 -/- :: Col6a1 -/ - mice showed a significantly increased deposition of fibrillar collagens in the endomysial ECM when compared to the corresponding WT samples. A similar increase was however observed in Col6a1 -/- mice ( Fig. 5D, E ). Thus, both in the absence and upon abnormal buildup of COL6, fibrillar collagens increase in muscle ECM, with no enhanced effect induced by ANTXR2 ablation. Altogether, these data indicate that the hypertrophic phenotype and the fibrillar collagen accumulation displayed by Antxr2 -/- muscles are not merely caused by the absence of ANTXR2 itself, nor the increase in fibrillar collagens, but by the progressive buildup of COL6. ANTXR2-deficient mice have impaired motor function Finally, we submitted mice to a battery of behavioural and physical tests. Importantly, Antxr2 -/- and age-matched WT mice had similar voluntary activity during both nocturnal and diurnal, at 3-and 8-months ( Fig. 6A and Supplementary Fig. 7A). To specifically evaluate muscle function, we determined plantar flexor muscle force by measuring torque production after electrical stimulation of the sciatic nerve in anaesthetized 8-month-old mice mice 40 . Of note, as for the muscles analysed above, the plantar flexor muscles of ANTXR2-deficient mice showed hypertrophy (Supplementary Fig. 7B). Surprisingly, at tetanus, the mice, whether Antxr2 -/- or WT, showed similar absolute or normalized force ( Fig. 6B and Supplementary Fig. 7C, D). Thus, the displayed muscle hypertrophy did not translate into an increase in strength. Interestingly, 8-month-old Antxr2 -/- mice showed enhanced maximal force transmission speed, particularly at lower stimulation frequencies (Supplementary Fig. 7E) and a trend toward slower relaxation time after a single twitch (Supplementary Fig. 7F, G). These altered parameters may result from the increased stiffness of ANTXR2-deficient muscles, which could reduce the in-series compliance of a contracting muscle. Download figure Open in new tab Figure 6. Antxr2 -/- mice display multiple signs of muscle weakness . ( A ) Quantification of locomotor activity of 8-month-old male WT and Antxr2 -/- mice, as determined by 24-h-long home cage monitoring. The histograms show the sum of total ambulatory and fine movements during the entire day (Total) or only during the diurnal (Light cycle) or nocturnal (Dark cycle) periods. Data are expressed as mean ± s.e.m. ( n = 11-12, each group; NS, not significant; two-tailed Mann-Whitney test). (C) Histograms showing the absolute and normalized force in plantar flexor muscles of 8-month-old WT and Antxr2 -/- male mice ( n = 19-29 muscles from 10 WT and 15 Antxr2 -/- mice, respectively; NS, not significant; *, P <0.05; ** P <0.01; ***, P <0.001; two-tailed Mann-Whitney test). ( C ) Quantification of the rotarod performance test of 8-month-old WT and Antxr2 -/- female and male mice. Histograms represent the mean latency time. Red dots indicate the animals that reached the maximum time (500 s) in at least one of the two measurements. Data are expressed as mean ± s.e.m. ( n = 15, each group; *, P <0.05; two-tailed Mann-Whitney test). ( D ) Quantification of the four-limb hanging performance of 8-month-old WT and Antxr2 -/- female and male mice. Histograms represent the holding impulse (i.e. best holding time normalized for body weight of the animal). Red dots indicate the animals that reached the maximum holding time (600 s) in at least one of the two evaluation tests. Data are expressed as mean ± s.e.m. ( n = 14–17, each group; *, P <0.05; ** P < 0.01; two-tailed Mann-Whitney test). ( E ) Quantification of the treadmill exercise performance of 8-month-old WT and Antxr2 -/- male mice. Histograms represent the time to fatigue (left panel) and the total distance run (right panel). Data are expressed as mean ± s.e.m. ( n = 20, each group; *, P <0.05; **, P <0.01; two-tailed Mann-Whitney test). We next investigated the motor coordination and tolerance to fatigue of 8-month-old male and female mice. Antxr2 -/- mice showed a significantly lower performance when subjected to rotarod ( Fig. 6C ) and four-limb hanging ( Fig. 6D ) tests, differences that were not seen in younger mice (Supplementary Fig. 7H, I). Then, mice were further subjected to a treadmill running protocol. At the age of 8-months, when forced to run until they reached exhaustion, Antxr2 -/- animals performed worse then age-matched WT animals, showing a decline in both the time needed to reach maximum exercise capacity and the total distance covered ( Fig. 6E ). This difference was not observed in younger mice, further emphasizing the progressive nature of the alterations induced by ANTRX2 deficiency (Supplementary Fig. 7J). Altogether, these results indicate that ANTXR2-deficient mice undergo a progressive impairment of locomotor activity and a decreased physical fitness during adult life. DISCUSSION Mutations in COL6 genes that impair COL6 deposition lead to myopathy, highlighting the essential role of this non-fibrillar collagen in maintaining the proper architecture and function of the extracellular matrix 24 . While the pathological consequences of COL6 deficiency in skeletal muscle have been well documented, less is known about the importance of tuning the proper abundance of COL6 in the ECM. The amount of a given protein in the ECM is the result of biogenesis and turnover, with the first depending largely on gene expression and the second on degradation. ECM protein turnover is thought to occur primarily through extracellular processing by matrix metalloproteases (MMPs), a class of secreted, broad-substrate, Zn 2+ -dependent endopeptidases 5 . An alternative route for ECM and collagen remodelling is however emerging, where degradation occurs intracellularly following receptor-mediated endocytosis and transport to lysosomes 9 , 10 . This route is particularly relevant to COL6 given its relative resistance to MMPs 41 – 44 , possibly due to its distinctive structure, characterized by a somewhat short triple-helical region and large globular domains. While several receptors have been found to bind COL6 17 , ANTXR2 appears unique in its ability to mediate cellular uptake of COL6 and targeting to lysosomes 11 . In this study, we therefore examined the role of ANTXR2 in skeletal muscle and found that it is predominantly expressed by interstitial FAPs, with negligible expression by mature myofibers. In vivo ANTXR2 ablation in Antxr2 -/- mice results in the early and progressive COL6 buildup in intramuscular ECM, leading to elevated amounts of fibrillar collagens and increased stiffness in adult mice. These ECM changes are accompanied by myofiber changes, with a non-functional hypertrophy and clearly identifiable signs of functional impairment and muscle weakness. These findings underline that proper control of extracellular COL6 levels — through both gene expression and ANTXR2-mediated degradation — is crucial for maintaining muscle function, as both deficiency and excess of COL6 are detrimental. Antxr2 -/- mice appear normal at birth and shortly after. At four weeks, they start a progressive accretion of extracellular COL6 in the intramuscular connective tissue. This indicates that, under physiological conditions, ANTXR2-mediated COL6 degradation needs to occur already in young mice to ensure proper homeostasis of this key muscle ECM component. The progressive COL6 buildup in muscles of Antxr2 -/- mice is paralleled by an ensuing accumulation of fibrillar collagens, which is particularly prominent at 8 months of age. In contrast, the basal lamina of ANTXR2-deficient muscles shows no apparent sign of alterations, indicating that defects are restrained to the outer part of the intramuscular connective tissue layers. In healthy muscle, the ECM is most synthesized and remodelled by FAPs 36 . Interestingly, both COL6 and ANTXR2 are abundantly expressed by FAPs, indicating that these cells both produce and degrade COL6. Of note, ablation of Antxr2 does not lead to upregulation of fibrotic markers, such as Acta2 and Lox , neither in muscles nor in FACS-sorted FAPs. Therefore, the progressive accumulation of COL6 does not trigger myofibroblast differentiation. Consistently, we did not detect other typical signs of fibrotic changes, such as fibrous-adipose replacement and cellular infiltration 32 . This is further supported by electron microscopy imaging, which, while revealing a pronounced accumulation of fibrillar collagens in the intramuscular connective tissue of adult Antxr2 -/- mice, does not show the chaotic and irregular arrangement of collagen fibrils typical of a fibrotic condition. Taken together, these data indicate that the lack of either COL6 or ANTXR2 leads to increased fibrillar collagens in the ECM, but the processes underlying such ECM changes involve distinct mechanisms. Both mice and patients with COL6 deficiency display upregulation of genes coding for COL1 and COL3, consistently with the presence of muscle fibrosis 45 , 46 . In the absence of ANTXR2, however, expression of collagen genes is unchanged, but COL6 accumulates due to altered degradation, and as a secondary effect fibrillar collagens accumulate in the ECM, most likely due to the ability of COL6 microfilaments to tightly interconnect with collagen fibrils 47 – 51 . Our data on the biomechanical properties of TA muscles from adult Antxr2 -/- mice indicate that the progressive accumulation of COL6 and fibrillar collagens in the ECM is paralleled by significant changes in the elastic modulus, with increased muscle stiffness. Interestingly, this also occurs in muscles of 3-month-old Antxr2 -/- animals, when COL6 – but not fibrillar collagens – already display increased ECM deposition. This observation supports earlier findings indicating that COL6 contributes to the fine regulation of tissue stiffness, affecting the mechanical properties of the tissue 47 , 48 , 52 , 53 . Although ANTXR2 expression by mature myofibers is very low, its ablation and the ensuing progressive accretion of COL6 in the ECM lead to a number of distinctive myofiber changes. First, myofibers of adult Antxr2 -/- male mice consistently show the presence of intracellular membranous structures positive for Gomori staining and with features of tubular aggregates. Such tubular aggregates represent myofiber inclusions consisting of tubules derived from the sarcoplasmic reticulum, typically more abundant in males, found in a variety of muscle disorders characterized by weakness, myalgia and cramps, being also described in muscles undergoing age-related sarcopenia 33 , 34 . The most remarkable change elicited by ANTXR2 ablation is however myofiber hypertrophy, associated with increased protein synthesis. This hypertrophic phenotype is linked to the progressive COL6 buildup, since muscles of Antxr2 -/- :: Col6a1 -/ - double knockout mice showed signs of myofiber atrophy, rather than hypertrophy, akin to what is displayed by muscles of Col6a1 -/- mice 40 . Notably, the myofiber hypertrophy displayed by ANTXR2-deficient muscles does not yield increased force, leading instead to functional defects, as revealed by various behavioural tests and in vivo quantification of muscle contractile force showing that Antxr2 -/- mice experience impaired locomotor activity and reduced physical fitness when physically challenged. Taking into account that myofibers of adult WT muscles barely express ANTXR2, these results strongly supports the concept that alterations in ANTXR2-deficient FAPs, in particular their inability to internalize COL6, result in ECM buildup in Antxr2 -/- muscles, in turn leading to increased muscle stiffness and compromised lateral transmission of force between myofibers and from muscle to tendons, especially during high-intensity exercise. In conclusion, our data indicate that ANTXR2 is required for the structural integrity and function of skeletal muscle. In particular, it contributes to the regulation of the extracellular abundance of COL6 and thereby to the proper organization and dynamics of the ECM. As such, ANTRX2 represents an ECM regulator that may have significant implications in various physiological and pathological contexts where ECM remodelling occurs, such as aging and fibrosis. Loss-of-function mutations of the human ANTXR2 gene lead to a devastating rare genetic disease, Hyaline Fibromatosis Syndrome (HFS) 11 , 15 . Consistent with the here identified role of ANTXR2 in muscle functions, HFS patients commonly experience muscle stiffness, weakness and joint contractures 54 . In particular, prominent skeletal muscle involvement with myopathic changes was reported in some infantile patients 55 , 56 . Thus, our findings indicate that Antxr2 -/- mice also represent a valuable model for investigating the pathomolecular mechanisms underlying muscle alterations in HFS. MATERIAL AND METHODS Mouse maintenance Age-matched WT, ANTXR2 knockout ( Antxr2 -/- ), COL6 knockout ( Col6a1 -/- ) 57 and ANTXR2/COL6 double knockout ( Antxr2 -/- :: Col6a1 -/- ) mice were maintained in the C57BL/6J background and housed in specific pathogen-free controlled environment (23° C temperature, 12 h light/12 h dark cycle). Mice were provided with ad libitum standard chow and water. Antxr2 -/- mice were generated by targeted deletion of exon 3 of the Antxr2 gene 11 , while Antxr2 -/- :: Col6a1 -/- double knockout mice were generated by crossing Antxr2 +/– and Col6a1 +/– heterozygous animals. According to the experiment, 5-day-old pups or 4-, 12-and 35-week-old male and female mice were used. All procedures were performed according to protocols approved by the Veterinary Authorities of the Canton Vaud (licences VD 2144.4 and VD 3918) and by the Ethics Committee of the University of Padova (OPBA, n. 100/2020-PR) and in agreement with the Swiss and Italian pertinent laws. Hindlimb muscle dissection Hindlimb skin and connective tissue (epimysium) were gently removed to expose the underlying tibialis anterior (TA), quadriceps femoris and gastrocnemius muscles. For 5-day-old pups, given the small size of skeletal muscles and the technical challenges of a proper tissue dissection, only the gastrocnemius muscle was collected and analyzed. Once harvested, muscles were quickly immersed in an isopentane-containing beaker, precooled by apposition to liquid nitrogen vapours. Frozen muscles were then stored at-80° C until subsequent analyses. Where indicated, extensor digitorum longus (EDL) muscles were collected and immediately fixed with 4% paraformaldehyde for 60 min at 4° C, washed twice in phosphate-buffered saline (PBS) and stored in PBS at 4° C. Before immunofluorescent analysis, EDL muscles underwent mechanical dissociation using scalpels to isolate either single myofibers or myofiber bundles. Histological analyses Cross-sections (10-μm-thick) of frozen muscles were used for the following staining protocols. For hematoxylin-eosin staining, slides were hydrated in bidistilled water for 2 min and stained with haematoxylin (Sigma-Aldrich) for 1-2 min, followed by washing with bidistilled water for 3 min. The slides were incubated for 1 min with eosin (Sigma-Aldrich), and subsequently rinsed in bidistilled water. For sirius red staining, slides were incubated in 4% paraformaldehyde for 15 min at room temperature, washed twice in PBS and incubated with 300 μL of picrosirius red solution (Sigma-Aldrich) in a moist chamber for 1 h at room temperature. Slides were then washed four times for 5 min in 0.5% glacial acetic acid (Sigma-Aldrich) under gentle shaking. For modified Gömöri trichrome staining, slides were hydrated in bidistilled water for 2 min and then incubated with Gomori trichrome (CliniSciences) for 10 min, and differentiated with 0.2% acetic acid for 10 min. After dehydration, slides were immersed in xylene and mounted with Eukitt Quick-hardening mounting medium (Sigma-Aldrich). Bright-field images were captured by using a Leica DM-R microscope equipped with a digital camera or an Olympus VS200 slide scanner. Sirius red staining images were acquired with a digital camera-equipped Leica DM6 B microscope under polarized light. Gomori-positive cytoplasmic inclusions were manually counted on the entire stained muscle section. Quantification of sirius red staining was performed with Fiji 58 . Immunofluorescence imaging Cross-sections (10-μm-thick) of frozen muscles were fixed and permeabilized in cold 1:1 methanol-acetone solution for 10 min at - 20° C, washed twice in PBS under gentle shaking and saturated with 10% goat serum (GS) in PBS at room temperature. Single myofibers or myofiber bundles were permeabilized with 0.2% Triton X-100 in 10% GS for 20 min and finally saturated with 10% GS in PBS at room temperature. After 60 min, primary antibodies were prepared in 5% GS and incubated overnight at 4° C. The following primary antibodies were used: rat monoclonal anti-ANTXR2 (1:50; clone 8F7, homemade produced 11 ); rabbit recombinant anti-Vimentin (1:200; D21H3, Cell-signaling technology); rabbit polyclonal anti-COL6 α3 (1:100; kindly provided by Dr. Raimund Wagener, Cologne 59 ); guinea pig polyclonal anti-COL6 α6 (1:200; kindly provided by Dr. Raimund Wagener, Cologne) 59 ; rabbit polyclonal anti-COL1 α1 (1:200; BP2201, Acris); rabbit polyclonal anti-COL3 α1 (1:200; 22734-1-AP, Proteintech); rabbit anti-Laminin (1:800; L9393, Sigma-Aldrich); rabbit polyclonal anti-FN1 (1:200; F3648, Sigma-Aldrich); rat monoclonal anti-HSPG2 (1:200; sc-33707, Santa Cruz Biotechnology); rabbit polyclonal anti-COL4 (1:200; AB756P, Millipore); mouse monoclonal anti-Myosin Heavy Chain IIA (1:100; SC-71, Developmental Studies Hybridoma Bank); rabbit polyclonal anti-Calsequestrin (1:200; PA1-913, Thermo-Fisher); mouse monoclonal anti-Stromal Interaction Molecule 1 (STIM1, 1:100; 610954, BD Transduction Laboratories); rabbit polyclonal anti-Calcium Release-Activated Calcium Modulator 1 (CRACM1/Orai1, 1:100; 4281, ProSci inc.); mouse monoclonal anti-Dihydropyridine Receptor (DHPR, 1:100; clone MA3-920, Sigma-Aldrich); mouse monoclonal anti-Sarco/Endoplasmic Reticulum Calcium ATPases (SERCA1/2/3, 1:100; sc-271669, Santa Cruz Biotechnology). The next day, slides or fiber bundles were washed three times in PBS for 10 min under gentle shaking and then incubated with appropriate secondary antibodies (Jackson ImmunoResearch) and nuclear dye Hoechst 33258 for 1 h at room temperature. To stain F-Actin, Phalloidin conjugated with Alexa Fluor 647 (Thermo-Fisher) was added together with secondary antibodies. Slides and myofiber bundles were washed three times in PBS for 10 min under gentle shaking at room temperature, and mounted in ProLong Glass Antifade Mountant (Invitrogen). Images were acquired with a Leica SP8 Stellaris laser-scanning confocal microscope. Quantification of the thresholded area was performed with Fiji. RNA in situ hybridization RNA in situ hybridizations were performed using the RNAscope Fluorescent Multiplex V2 Assay (Bio-Techne, cat. n. 323110) following the manufacturer’s instructions. Briefly, 10-μm-thick cross-sections of TA were fixed with 4% paraformaldehyde for 60 min at 4° C, washed twice with PBS, dehydrated and quenched with hydrogen peroxide for 10 min at room temperature. Then, slides were washed twice in PBS and treated for 10 min at room temperature with protease IV. Probes for mouse Antxr2 (Bio-Techne, cat. n. 468651) were incubated for 2 h and RNAscope amplification was performed following the manufacturer’s instructions using Opal 650 fluorophore reagent pack (Akoya Biosciences, cat. n. FP1496001KT). Coupled immunofluorescence analysis was performed as described above without the permeabilization step. Finally, slides were mounted in Prolong Gold antifade mounting medium (Thermo-Fisher) and imaged using a Leica SP8 laser-scanning confocal microscope. Quantification of mRNA dots was performed with QuPath 60 . Muscle morphometric analysis Cross-sections (10-μm-thick) of frozen TA muscles were incubated with 1 μg/mL of wheat germ agglutinin (WGA) conjugated with Alexa Fluor 488 (Invitrogen) and 2.5 μg/mL of Hoechst 33258 (Sigma-Aldrich) in PBS for 20 min at room temperature. Slides were washed twice in PBS for 5 min under gentle shaking and mounted in 80% glycerol in PBS. For each sample, the entire stained section was acquired with a Leica SP8 Stellaris laser-scanning confocal microscope (Leica Microsystems). Myofiber cross-sectional area (CSA) and minimum Feret diameter (MFD) were measured with the Myosoft plugin 61 . Transmission electron microscopy TA muscles were fixed overnight at 4° C with 2.5% glutaraldehyde in 0.1 M cacodylate buffer, washed with 0.1 M cacodylate buffer, post-fixed for 2 h with 1% osmium tetroxide, and embedded in Epon812 (Electron Microscopy Sciences). Transversal and longitudinal ultrathin sections were stained with uranyl acetate and lead citrate and observed with a Philips EM400 electron microscope operating at 100 kV. Western blotting Frozen muscles were pulverized by grinding in liquid nitrogen and lysed in extraction buffer (50 mM Tris-HCl, pH 7.5; 150 mM NaCl; 10 mM MgCl 2 ; 1 mM EDTA; 10% glycerol; 0.5 mM DTT; 2% SDS; 1% Triton X-100) supplemented with protease and phosphatase inhibitors (Sigma-Aldrich). SDS-insoluble fractions were obtained after centrifuging the protein extracts and lysing the pellet in urea buffer (8 M urea, 2% SDS) supplemented with protease inhibitors (Sigma-Aldrich). Protein concentration was determined by the BCA Protein Assay Kit (Thermo-Fisher) and 30-40 μg of protein samples were used for SDS-PAGE in polyacrylamide Novex NuPAGE Bis-Tris 4-12% gels (Invitrogen). For native COL6 detection, protein samples were run in a homemade composite 2.5% acrylamide/0.5% agarose gel under non-reducing conditions for higher molecular weight separations 62 . Gels were electrotransferred onto polyvinylidene difluoride membranes (Millipore), stained with Ponceau S (Sigma-Aldrich) and blocked for 1 h with 5% non-fat dry milk in Tris-buffered saline solution containing 0.1% Tween 20 (TBS-T), followed by overnight incubation at 4° C with primary antibodies. The following primary antibodies were used: rabbity polyclonal anti-COL6 (1:1,000; 70-XR95, Fitzgerald Industries, for native COL6 detection); rabbit polyclonal anti-COL6 α1 (1:1,000; sc-20649, Santa Cruz Biotechnology); rabbit polyclonal anti-COL6 α3 (1:1,000; homemade produced, kindly provided by Dr. Raimund Wagener, Cologne) 59 ; rabbit polyclonal anti-COL3 α1 (1:2,000; 22734-1-AP, Proteintech); mouse monoclonal anti-GAPDH (1:125,000; MAB374, Millipore); mouse monoclonal anti-Vinculin (1:1,000; V4505, Sigma-Aldrich); mouse monoclonal anti-puromycin (1:2,500; clone 12D10, Millipore). After three washes in TBS-T, membranes were incubated for 1 h with horseradish peroxidase-conjugated anti-rabbit or anti-mouse secondary antibodies (1:1,000; Bethyl Laboratories). Detection was carried out by SuperSignal West Pico (Thermo-Fisher) and densitometric quantification was performed by Fiji software. Hydroxyproline assay For this assay the gastrocnemius muscle was selected based on its larger size, enabling the collection of sufficient tissue material required for the analyses. Muscles were pulverized, collected in a pre-cooled tube, weighted, and maintained over liquid nitrogen vapours. Next, 7 μL of HCl and 7 μL of bidistilled water were added for each mg of tissue, and the tubes were kept at 130° C for 3 h to dissolve the tissue sample. Lysates were further homogenized by pushing the specimens through a 26G needle four times and then centrifuged at 12,000 rcf for 10 min at room temperature. 30 μL of supernatant were transferred into a new tube and 625 μL of chloramine T buffer (Sigma-Aldrich) were added to each sample and incubated for 20 min at room temperature. Then, 625 μL of 4-(dimethylamino)benzaldehyde (DMAB) buffer (Sigma-Aldrich) was added and incubated for 10 min at 65° C. Reaction was stopped by putting tubes on ice and 200 μL of each sample were loaded in duplicate in a 96-well plate. Color intensity was measured with a microplate spectrophotometer reader (Tecan Life Sciences), by exciting the samples at a wavelength of 540 nm. A calibration curve obtained from known concentrations of hydroxyproline (Sigma-Aldrich) was used to extrapolate the concentrations of the samples from their experimental absorbances. In vivo protein synthesis In vivo protein synthesis in skeletal muscle was measured using the SUrface SEnsing of Translation (SUnSET) technique 39 . Mice were weighed and subjected to injection with puromycin (Sigma-Aldrich) dissolved in 100 μL of sterile PBS at the concentration of 0.04 μmol/g body mass. 30 min after puromycin injection, TA muscles were collected, frozen in liquid nitrogen and stored (−80 °C) for western blot analysis. Fluorescence-activated cell sorting FAPs were obtained from gastrocnemius muscle using a previously described protocol with slight modifications 63 . In brief, muscle tissue underwent enzymatic digestion with 2 mg/mL Collagenase II (Sigma-Aldrich), 1.5 U/mL Collagenase D (Sigma-Aldrich), 2.4 U/mL Dispase II (Sigma-Aldrich), and 250 mM CaCl 2 . Following digestion, muscles were gently mashed and supplemented with red blood cell lysis buffer (0.15 mM NH 4 Cl, 8.4 mM KHCO 3 , 1.2 mM EDTA in PBS) before filtration through 70 µM and 40 µM cell strainers to obtain single-cell suspensions. These suspensions were then incubated with primary antibodies targeting specific cell markers for 30 min at 4° C in a rotating wheel. The following antibodies were used: CD45-eFluor 450 (1:50; Invitrogen, clone 30-F11), CD31 (PECAM-1)-PE-Cy7 (1:50; Invitrogen, clone 390), CD140a (PDGFRa)-APC (1:25; Invitrogen, clone APA5), Ly6A/E (Sca-1)-FITC (1:50; Invitrogen, clone D7), Viability Fixable Dye 405/520 (1:100; Miltenyi Biotec). FAPs (CD45 - /CD31 - /PDGFRα + /Sca-1 + ) were subsequently isolated using flow sorting in a FACS Canto II (Becton Dickinson). Sorted FAPs were collected into cold PBS before RNA extraction. RT-qPCR Muscle samples were pulverized by grinding in liquid nitrogen and total RNA was extracted using 0.5 mL TRIzol Reagent (Thermo-Fisher). RNA extraction from sorted FAPs was performed using the Nucleospin RNA XS isolation kit (Macherey-Nagel). RNA was quantified with Nanodrop ND-1000 (Nanodrop Technologies). Reverse transcription was performed with 500 ng total RNA and M-MLV Reverse Transcriptase (Thermo-Fisher), using random hexamers as previously described 64 . The resulting cDNAs were processed for quantitative real-time PCR using Rotor-Gene SYBR Green PCR Kit mastermix (Qiagen) and a Rotor-GeneQ thermocycler (Qiagen). Each sample was loaded in triplicate and analyzed with RotorGene Q 2.0.24 software. Rps16 or Gapdh were used as housekeeping gene controls. Primers are listed in Supplementary Table 1. Quantification of muscle stiffness The mechanical properties of fresh TA muscles were characterized through a tensile test using a uniaxial testing machine (ZwickRoell ProLine Z005) equipped with a 10 N load cell. Muscle samples were fixed to the clamps by using two suture needles with inextensible thread, suitably shaped to prevent the specimen from slipping. The test was performed by imposing a strain rate of 10% of the initial length (distance between the needles) per minute, and was stopped at sample failure, acquiring total displacement (mm) and standard force (N). Data were then analyzed in MS Excel ® , obtaining strain (%) and stress (kPa) values and plotting the stress-strain curves. The elastic modulus was then determined by performing a linear regression on the linear portion of the curves. Spontaneous locomotor activity Global tracking of the voluntary activity over 24 h, covering both nocturnal and diurnal periods was measured with a Comprehensive Laboratory Animal Monitoring System (CLAMS, Columbus Instruments). Mice were placed individually in the plexiglas chambers and the spontaneous activity, the food intake and the water intake were measured simultaneously. Ambulatory movements and fine movements were measured. Four-limb hanging test Mice were placed on a grid, which was kept upside down above an empty cage filled with bedding materials. The hanging time, i.e., the time until the animal fell, was recorded. A fixed limit of 900 s was set, and each mouse was tested twice with a 30-min break between measurements. Holding impulse was obtained by multiplying the hanging time for the respective mouse body weight. The maximum holding time was used for the analysis according to TREAT-NMD standard operating procedures (DMD_M.2.1.005). Rotarod performance test Two days before the experiments, mice underwent training sessions on a rotating rod (Ugo Basile) for two sessions lasting 10 min each, maintained at a constant speed of 7 rpm, with a 30-min break in between sessions. The day of the experiment, after 3 min warmup at a constant speed of 7 rpm, a ramp of 7-30 rpm in 300 s was initiated. The ending time point was set at 500 s, and mice that did not fall off within this period were given a maximum score. The test was repeated after 30 min and the average value was used for statistics. Exercise performance test Before the test, mice were habituated to the treadmill (Panlab) for 10 min at a speed of 5 cm/s. Acute exercise was started at the very moderate speed of 15 cm/s with a 10° positive inclination, and the speed was gradually increased of 3 cm/s every 12 min. The run distance and the number of received shocks (0.1-0.3 mA) were monitored. If a mouse received more than five shocks in two consecutive minutes, it was considered as having reached exhaustion and placed back in its homecage. In vivo muscle force measurement Mice were anesthetized with one i.p. injection of a solution containing 100 mg/kg ketamine and 10mg/kg xylazine and, a few minutes later, one i.p. injection of 5 mg/kg altadol as a pain relief treatment. A small incision was made from the knee to the hip and the sciatic nerve was exposed. Teflon-coated multistranded stainless-steel wires (AS 632; Cooner Wire Company, Chatsworth, CA, USA) was sutured with 5-0 silk thread at each side of the nerve and the wound was closed using the same thread. To avoid recruitment of the ankle dorsal flexors, and the consequential reduction of torque, the common peroneal nerve was cut. Torque production of the stimulated plantar flexors was measured using a muscle lever system (Model 305C; Aurora Scientific, Aurora, ON, Canada), data were acquired using the DMCv5.3 software (Aurora Scientific), and electrical stimulation was achieved using a Grass S11 stimulator (Grass Instrument Company, Quincy, MA, USA). The force-frequency curves were determined by stepwise increasing stimulation frequency (from single pulse to 20, 40, 55, 75, 100 and 150 Hz; train duration, 200 ms; single pulse width 500 µsec; 4-6 V), with a pause of 30 s between stimuli to avoid muscle fatigue. Time parameters (time to peak and half relaxation time) were measured on the force trace obtained with a single stimulus (twitch). Contraction parameters were obtained from the tetanus trace obtained at 100 Hz electrical stimulation. The fast rate of force development was measured as a linear fitting (max speed, g/ms). Normalized force was obtained by dividing absolute force to muscle weight (obtaining kN/g). Representative diagrams of twitch and absolute force traces are shown in Fig. S7G . Raw data analysis was performed using a custom-made automatic LabView software. Single-nucleus RNA sequencing dataset analysis The GSE147127 dataset, containing single-nucleus RNA-seq data of skeletal muscles from C57BL/6 mice of different ages 28 , was used for this analysis. Normalized count matrix and dimensional reduction data were downloaded from https://www.synapse.org/#!Synapse:syn21676145/files/ as Seurat object. Seurat v4.3.0.1 was then used for plotting features of interest on the pre-computed UMAP 65 . The dotPlot function from scCustomize v1.1.3 was used to plot the percentage of cells expressing selected genes. Statistics All results are provided as mean ± s.e.m. Statistical analysis of data was performed with an unpaired two-tailed Mann-Whitney test (GraphPad). For experiments with more than two conditions, one-way or two-way analysis of variants (ANOVA) was used (GraphPad). When ANOVA revealed significant differences, further analysis was carried out using Dunnet’s or Sidak’s multiple comparison test. Statistical significance was set at P <0.05. All histograms display individual values, and the number of biological replicates (always greater than three) as well as the specific statistical tests used are indicated in the figures’ captions. Author contributions S.M., F.G.v.d.G. and P.B. designed the study. S.M. performed most of the experiments and relative analyses. L.J.B., M.S., L.A., B.K., and L.G. contributed to in vivo experiments. J.D. analyzed transcriptomic data. L.N. and B.B. performed in vivo muscle force measurements. G.P. and N.F. contributed to hydroxyproline content analysis. P.S. performed transmission electron microscopy analysis. A.D.A., G.M.F., P.Br. and G.V. performed muscle stiffness measurements. S.M., F.G.v.d.G. and P.B. wrote the original draft and all the other authors contributed to the final version of the manuscript. Acknowledgements We thank Dr. Dario Bizzotto (University of Padova) and Dr. Audrey Chuat (EPFL) for the maintenance of mouse colonies and genotyping; Dr. Olha Novokhatska (EPFL) for the initial involvement in the locomotor activity tracking; Dr. Raimund Wagener (University of Cologne, Cologne, Germany) for providing COL6 antibodies; the core facilities BIOP and CPG at EPFL for sharing their equipment and expertise; Dr. Matilde Cescon (University of Padova) and all the members of G. van der Goot and P. Bonaldo labs for helpful discussions and suggestions. This work was funded by the Italian Ministry of University and Research (Grant 2022MXH3JY to P.B.), the Telethon Foundation (GGP19229 to P.B.), and the Swiss Foundation for Research on Muscle Diseases to F.G.v.d.G.. S.M. was supported by the European Molecular Biology Organization EMBO Scientific Exchange (Grant Number 9111) and by the Marie-Curie MSCA Global Fellowship (Grant agreement 101149789). REFERENCES 1. ↵ Frantz , C. , Stewart , K. M. & Weaver , V. M . The extracellular matrix at a glance . J Cell Sci 123 , 4195 – 4200 ( 2010 ). OpenUrl FREE Full Text 2. ↵ Hynes , R. O. & Naba , A . Overview of the matrisome - An inventory of extracellular matrix constituents and Functions . Cold Spring Harb Perspect Biol 4 , a004903 ( 2012 ). 3. ↵ Cox , T. R. & Erler , J. T . Remodeling and homeostasis of the extracellular matrix: implications for fibrotic diseases and cancer . Dis Model Mech 4 , 165 – 178 ( 2011 ). OpenUrl Abstract / FREE Full Text 4. Maldonado , M. & Nam , J . The role of changes in extracellular matrix of cartilage in the presence of inflammation on the pathology of osteoarthritis . Biomed Res Int 2013, 284873 ( 2013 ). 5. ↵ Naba , A . Mechanisms of assembly and remodelling of the extracellular matrix . Nat Rev Mol Cell Biol 25 , 865 – 885 ( 2024 ). OpenUrl CrossRef PubMed 6. ↵ Winkler , J. , Abisoye-Ogunniyan , A. , Metcalf , K. J. & Werb , Z . Concepts of extracellular matrix remodelling in tumour progression and metastasis . Nat Commun 11 , 5120 ( 2020 ). 7. ↵ Chung , L. et al. Collagenase unwinds triple-helical collagen prior to peptide bond hydrolysis . EMBO J 23 , 3020 – 3030 ( 2004 ). OpenUrl Abstract / FREE Full Text 8. ↵ Manka , S. W. et al. Structural insights into triple-helical collagen cleavage by matrix metalloproteinase 1 . Proc Natl Acad Sci U S A 109 , 12461 – 12466 ( 2012 ). OpenUrl Abstract / FREE Full Text 9. ↵ Rainero , E . Extracellular matrix endocytosis in controlling matrix turnover and beyond: emerging roles in cancer . Biochem Soc Trans 44 , 1347 – 1354 ( 2016 ). OpenUrl Abstract / FREE Full Text 10. ↵ Wagenaar-Miller , R. A. et al. Complementary roles of intracellular and pericellular collagen degradation pathways in vivo . Mol Cell Biol 27 , 6309 – 6322 ( 2007 ). OpenUrl Abstract / FREE Full Text 11. ↵ Bürgi , J. et al. CMG2/ANTXR2 regulates extracellular collagen VI which accumulates in hyaline fibromatosis syndrome . Nat Commun 8 , 15861 ( 2017 ). 12. ↵ Bürgi , J. et al. Ligand binding to the Collagen VI receptor triggers a Talin-to-RhoA switch that regulates receptor endocytosis . Dev Cell 53 , 418 – 430 ( 2020 ). OpenUrl CrossRef PubMed 13. ↵ Liu , S. et al. Capillary morphogenesis protein-2 is the major receptor mediating lethality of anthrax toxin in vivo . Proc Natl Acad Sci U S A 106 , 12424 – 12429 ( 2009 ). OpenUrl Abstract / FREE Full Text 14. ↵ Sergeeva , O. A. & van der Goot , F. G . Converging physiological roles of the anthrax toxin receptors . F1000Res 8 , 1415 ( 2019 ). OpenUrl 15. ↵ Dowling , O. et al. Mutations in capillary morphogenesis gene-2 result in the allelic disorders juvenile hyaline fibromatosis and infantile systemic hyalinosis . Am J Hum Genet 73 , 957 – 966 ( 2003 ). OpenUrl CrossRef PubMed Web of Science 16. ↵ Peters , D. E. et al. Capillary morphogenesis protein-2 is required for mouse parturition by maintaining uterine collagen homeostasis . Biochem Biophys Res Commun 422 , 393 – 397 ( 2012 ). OpenUrl CrossRef PubMed 17. ↵ Cescon , M. , Gattazzo , F. , Chen , P. & Bonaldo , P . Collagen VI at a glance . J Cell Sci 128 , 3525 – 3531 ( 2015 ). OpenUrl Abstract / FREE Full Text 18. ↵ Bonaldo , P. , Russo , V. , Bucciotti , F. , Doliana , R. & Colombatti , A . Structural and functional features of the alpha3 chain indicate a bridging role for chicken collagen VI in connective tissues . Biochemistry 29 , 1245 – 1254 ( 1990 ). OpenUrl CrossRef PubMed 19. ↵ Tillet , E. et al. Recombinant expression and structural and binding properties of α1(VI) and α2(VI) chains of human collagen type VI . Eur J Biochem 221 , 177 – 187 ( 1994 ). OpenUrl PubMed Web of Science 20. ↵ Sabatelli , P. et al. Collagen VI deficiency affects the organization of fibronectin in the extracellular matrix of cultured fibroblasts . Matrix Biol 20 , 475 – 486 ( 2001 ). OpenUrl CrossRef PubMed Web of Science 21. ↵ Wiberg , C. et al. Biglycan and decorin bind close to the n-terminal region of the collagen VI triple helix . J Biol Chem 276 , 18947 – 18952 ( 2001 ). OpenUrl Abstract / FREE Full Text 22. ↵ Jöbsis , G. J. et al. Type VI collagen mutations in Bethlem myopathy, an autosomal dominant myopathy with contractures . Nat Genet 14 , 113 – 115 ( 1996 ). OpenUrl CrossRef PubMed Web of Science 23. Camacho Vanegas , O. , et al. Ullrich scleroatonic muscular dystrophy is caused by recessive mutations in collagen type VI . Proc Natl Acad Sci U S A 98 , 7516 – 7521 ( 2001 ). OpenUrl Abstract / FREE Full Text 24. ↵ Bönnemann , C. G . The collagen VI-related myopathies: muscle meets its matrix . Nat Rev Neurol 7 , 379 – 390 ( 2011 ). OpenUrl CrossRef PubMed 25. ↵ Quarto , R. , Dozin , B. , Bonaldo , P. , Cancedda , R. & Colombatti , A . Type VI collagen expression is upregulated in the early events of chondrocyte differentiation . Development 117 , 245 – 251 ( 1993 ). OpenUrl Abstract / FREE Full Text 26. Braghetta , P. et al. Distinct regions control transcriptional activation of the alpha1(VI) collagen promoter in different tissues of transgenic mice . J Cell Biol 135 , 1163 – 1177 ( 1996 ). OpenUrl Abstract / FREE Full Text 27. ↵ Braghetta , P. et al. An enhancer required for transcription of the Col6a1 gene in muscle connective tissue is induced by signals released from muscle cells . Exp Cell Res 314 , 3508 – 3518 ( 2008 ). OpenUrl CrossRef PubMed Web of Science 28. ↵ Petrany , M. J. et al. Single-nucleus RNA-seq identifies transcriptional heterogeneity in multinucleated skeletal myofibers . Nat Commun 11 , 6374 ( 2020 ). 29. ↵ Zou , Y. , Zhang , R.-Z. , Sabatelli , P. , Chu , M.-L. & Bönnemann , C. G . Muscle interstitial fibroblasts are the main source of collagen VI synthesis in skeletal muscle: implications for congenital muscular dystrophy types Ullrich and Bethlem . J Neuropathol Exp Neurol 67 , 144 – 154 ( 2008 ). OpenUrl CrossRef PubMed 30. ↵ Gara , S. K. et al. Differential and restricted expression of novel collagen VI chains in mouse . Matrix Biol 30 , 248 – 257 ( 2011 ). OpenUrl CrossRef PubMed 31. ↵ Sabatelli , P. et al. Expression of collagen VI α5 and α6 chains in human muscle and in Duchenne muscular dystrophy-related muscle fibrosis . Matrix Biol 31 , 187 – 196 ( 2012 ). OpenUrl CrossRef PubMed Web of Science 32. ↵ Mann , C. J. et al. Aberrant repair and fibrosis development in skeletal muscle . Skelet Muscle 1 , 21 ( 2011 ). 33. ↵ Morgan-Hughes , J. A . Tubular aggregates in skeletal muscle: their functional significance and mechanisms of pathogenesis . Curr Opin Neurol 11 , 439 – 442 ( 1998 ). OpenUrl CrossRef PubMed Web of Science 34. ↵ Schiaffino , S . Tubular aggregates in skeletal muscle: just a special type of protein aggregates? Neuromuscul Disord 22 , 199 – 207 ( 2012 ). OpenUrl CrossRef PubMed 35. ↵ Sanes , J. R . The basement membrane/basal lamina of skeletal muscle . J Biol Chem 278 , 12601 – 12604 ( 2003 ). OpenUrl FREE Full Text 36. ↵ Gillies , A. R. & Lieber , R. L . Structure and function of the skeletal muscle extracellular matrix . Muscle Nerve 44 , 318 – 331 ( 2011 ). OpenUrl CrossRef PubMed Web of Science 37. ↵ Dooling , L. J. , Saini , K. , Anlaş , A. A. & Discher , D. E . Tissue mechanics coevolves with fibrillar matrisomes in healthy and fibrotic tissues . Matrix Biol 111 , 153 – 188 ( 2022 ). OpenUrl CrossRef PubMed 38. ↵ Schiaffino , S. & Reggiani , C . Fiber types in mammalian skeletal muscles . Physiol Rev 91 , 1447 – 1531 ( 2011 ). OpenUrl CrossRef PubMed 39. ↵ Schmidt , E. K. , Clavarino , G. , Ceppi , M. & Pierre , P . SUnSET, a nonradioactive method to monitor protein synthesis . Nat Methods 6 , 275 – 277 ( 2009 ). OpenUrl CrossRef PubMed Web of Science 40. ↵ Gambarotto , L. et al. Sustained oral spermidine supplementation rescues functional and structural defects in COL6-deficient myopathic mice . Autophagy 19 , 3221 – 3229 ( 2023 ). OpenUrl PubMed 41. ↵ Hey , S. & Linder , S . Matrix metalloproteinases at a glance . J Cell Sci 137 , jcs261898 ( 2024 ). 42. Myint , E. , Brown , D. J. , Ljubimov , A. V. , Kyaw , M. & Kenney , M. C . Cleavage of human corneal type VI collagen alpha 3 chain by matrix metalloproteinase-2 . Cornea 15 , 490 – 496 ( 1996 ). OpenUrl PubMed Web of Science 43. Motrescu , E. R. et al. Matrix metalloproteinase-11/stromelysin-3 exhibits collagenolytic function against collagen VI under normal and malignant conditions . Oncogene 27 , 6347 – 6355 ( 2008 ). OpenUrl CrossRef PubMed Web of Science 44. ↵ Kielty , C. M. , Lees , M. , Shuttleworth , C. A. & Woolley , D . Catabolism of intact type VI collagen microfibrils: susceptibility to degradation by serine proteinases . Biochem Biophys Res Commun 191 , 1230 – 1236 ( 1993 ). OpenUrl CrossRef PubMed 45. ↵ Guadagnin , E. et al. Transcriptome analysis of collagen VI-related muscular dystrophy muscle biopsies . Ann Clin Transl Neurol 8 , 2184 – 2198 ( 2021 ). OpenUrl PubMed 46. ↵ Mohassel , P. et al. Collagen type VI regulates TGF-β bioavailability in skeletal muscle in mice . J Clin Invest 135 , ( 2025 ). 47. ↵ Izu , Y. et al. Dysfunctional tendon collagen fibrillogenesis in collagen VI null mice . Matrix Biol 30 , 53 – 61 ( 2011 ). OpenUrl CrossRef PubMed Web of Science 48. ↵ Leiphart , R. J. et al. Coordinate roles for collagen VI and biglycan in regulating tendon collagen fibril structure and function . Matrix Biol Plus 13 , 100099 ( 2022 ). 49. Keene , D. R. , Engvall , E. & Glanville , R. W . Ultrastructure of type VI collagen in human skin and cartilage suggests an anchoring function for this filamentous network . J Cell Biol 107 , 1995 – 2006 ( 1988 ). OpenUrl Abstract / FREE Full Text 50. Keene , D. R. , Ridgway , C. C. & Iozzo , R. V . Type VI microfilaments interact with a specific region of banded collagen fibrils in skin . J Histochem Cytochem . 46 , 215 – 220 ( 1998 ). OpenUrl CrossRef PubMed 51. ↵ Bruns , R. R. , Press , W. , Engvall , E. , Timpl , R. & Gross , J. Type VI collagen in extracellular, 100-nm periodic filaments and fibrils: identification by immunoelectron microscopy . J Cell Biol 103 , 393 – 404 ( 1986 ). OpenUrl Abstract / FREE Full Text 52. ↵ Urciuolo , A. et al. Collagen VI regulates satellite cell self-renewal and muscle regeneration . Nat Commun 4 , 1964 ( 2013 ). 53. ↵ Zelenski , N. A. et al. Collagen VI regulates pericellular matrix properties, chondrocyte swelling, and mechanotransduction in articular cartilage . Arthritis Rheumatol 67 , 1286 – 1294 ( 2015 ). OpenUrl PubMed 54. ↵ Casas-Alba , D. et al. Hyaline fibromatosis syndrome: Clinical update and phenotype–genotype correlations . Hum Mutat 39 , 1752 – 1763 ( 2018 ). OpenUrl CrossRef PubMed 55. ↵ Zolkipli , Z. et al. Skeletal muscle involvement in infantile systemic hyalinosis . Eur J Paediatr Neurol 7 , 401 – 406 ( 2003 ). OpenUrl PubMed 56. ↵ Bayram , E. et al. Myopathic changes in muscle biopsy of a patient with Infantile Systemic Hyalinosis . Paediatr Croat 56 , 301 – 4 ( 2012 ). OpenUrl 57. ↵ Bonaldo , P. et al. Collagen VI deficiency induces early onset myopathy in the mouse: an animal model for Bethlem myopathy . Hum Mol Genet 7 , 2135 – 2140 ( 1998 ). OpenUrl CrossRef PubMed Web of Science 58. ↵ Schindelin , J. , et al. Fiji: an open-source platform for biological-image analysis . Nat Methods 9 , 676 – 682 ( 2012 ). OpenUrl CrossRef PubMed Web of Science 59. ↵ Gara , S. K. et al. Three novel collagen VI chains with high homology to the alpha3 chain . J Biol Chem 283 , 10658 – 10670 ( 2008 ). OpenUrl Abstract / FREE Full Text 60. ↵ Bankhead , P. et al. QuPath: Open source software for digital pathology image analysis . Sci Rep 7 , 16878 ( 2017 ). 61. ↵ Encarnacion-Rivera , L. , Foltz , S. , Hartzell , H. C. & Choo , H . Myosoft: An automated muscle histology analysis tool using machine learning algorithm utilizing FIJI/ImageJ software . PLoS One 15 , e0229041 ( 2020 ). OpenUrl CrossRef PubMed 62. ↵ Gualandi , F. et al. Autosomal recessive Bethlem myopathy . Neurology 73 , 1883 – 1891 ( 2009 ). OpenUrl CrossRef PubMed 63. ↵ Soliman , H. et al. Pathogenic potential of Hic1 -expressing cardiac stromal progenitors . Cell Stem Cell 26 , 205 – 220 ( 2020 ). OpenUrl CrossRef PubMed 64. ↵ Metti , S. , Da Ros , F. , Toniato , G. , Cescon , M. & Bonaldo , P . Native collagen VI delays muscle stem cell early differentiation . J Cell Sci 137 , jcs.261419 ( 2024 ). 65. ↵ Hao , Y. et al. Integrated analysis of multimodal single-cell data . Cell 184 , 3573 – 3587 .e29 ( 2021 ). OpenUrl CrossRef PubMed 66. Marsh , S. , Salmon , M. & Hoffman , P. samuel-marsh/scCustomize: Version 2.1.2 . Zenodo doi: 10.5281/zenodo.10724532 ( 2024 ). OpenUrl CrossRef View the discussion thread. Back to top Previous Next Posted September 19, 2025. Download PDF Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. You are going to email the following Antxr2-mediated fine-tuning of Collagen VI ensures skeletal muscle function Message Subject (Your Name) has forwarded a page to you from bioRxiv Message Body (Your Name) thought you would like to see this page from the bioRxiv website. Your Personal Message CAPTCHA This question is for testing whether or not you are a human visitor and to prevent automated spam submissions. Share Antxr2-mediated fine-tuning of Collagen VI ensures skeletal muscle function Samuele Metti , Lucie Josepha Bracq , Matteo Signor , Nicola Facchinello , Lisa Gambarotto , Leonardo Nogara , Giulia Pigato , Laurence Abrami , Béatrice Kunz , Julien Duc , Patrizia Sabatelli , Aurora De Acutis , Gabriele Maria Fortunato , Giovanni Vozzi , Paola Braghetta , Bert Blaauw , F. Gisou van der Goot , Paolo Bonaldo bioRxiv 2025.09.11.675515; doi: https://doi.org/10.1101/2025.09.11.675515 Share This Article: Copy Citation Tools Antxr2-mediated fine-tuning of Collagen VI ensures skeletal muscle function Samuele Metti , Lucie Josepha Bracq , Matteo Signor , Nicola Facchinello , Lisa Gambarotto , Leonardo Nogara , Giulia Pigato , Laurence Abrami , Béatrice Kunz , Julien Duc , Patrizia Sabatelli , Aurora De Acutis , Gabriele Maria Fortunato , Giovanni Vozzi , Paola Braghetta , Bert Blaauw , F. Gisou van der Goot , Paolo Bonaldo bioRxiv 2025.09.11.675515; doi: https://doi.org/10.1101/2025.09.11.675515 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Cell Biology Subject Areas All Articles Animal Behavior and Cognition (7642) Biochemistry (17715) Bioengineering (13907) Bioinformatics (42003) Biophysics (21470) Cancer Biology (18624) Cell Biology (25533) Clinical Trials (138) Developmental Biology (13390) Ecology (19935) Epidemiology (2067) Evolutionary Biology (24356) Genetics (15617) Genomics (22529) Immunology (17753) Microbiology (40432) Molecular Biology (17200) Neuroscience (88681) Paleontology (667) Pathology (2840) Pharmacology and Toxicology (4828) Physiology (7653) Plant Biology (15161) Scientific Communication and Education (2046) Synthetic Biology (4304) Systems Biology (9826) Zoology (2271)