Skip to main content Accessibility help
Hostname: page-component-cf9d5c678-5wlnc Total loading time: 1.371 Render date: 2021-08-03T04:47:44.064Z Has data issue: true Feature Flags: { "shouldUseShareProductTool": true, "shouldUseHypothesis": true, "isUnsiloEnabled": true, "metricsAbstractViews": false, "figures": true, "newCiteModal": false, "newCitedByModal": true, "newEcommerce": true, "newUsageEvents": true }

Invited review: mesenchymal progenitor cells in intramuscular connective tissue development

Published online by Cambridge University Press:  09 September 2015

Z. G. Miao
School of Animal Sciences, Henan Institute of Science and Technology, Xinxiang 453003, China Department of Animal Sciences, Washington State University, Pullman, WA 99164, USA
L. P. Zhang
Department of Animal Sciences, Washington State University, Pullman, WA 99164, USA Institute of Animal Science, Chinese Academy of Agricultural Sciences, Beijing 100193, China
X. Fu
Department of Animal Sciences, Washington State University, Pullman, WA 99164, USA
Q. Y. Yang
Department of Animal Sciences, Washington State University, Pullman, WA 99164, USA
M. J. Zhu
School of Food Science, Washington State University, Pullman, WA 99164, USA
M. V. Dodson
Department of Animal Sciences, Washington State University, Pullman, WA 99164, USA
M. Du
Department of Animal Sciences, Washington State University, Pullman, WA 99164, USA
E-mail address:


The abundance and cross-linking of intramuscular connective tissue contributes to the background toughness of meat, and is thus undesirable. Connective tissue is mainly synthesized by intramuscular fibroblasts. Myocytes, adipocytes and fibroblasts are derived from a common pool of progenitor cells during the early embryonic development. It appears that multipotent mesenchymal stem cells first diverge into either myogenic or non-myogenic lineages; non-myogenic mesenchymal progenitors then develop into the stromal-vascular fraction of skeletal muscle wherein adipocytes, fibroblasts and derived mesenchymal progenitors reside. Because non-myogenic mesenchymal progenitors mainly undergo adipogenic or fibrogenic differentiation during muscle development, strengthening progenitor proliferation enhances the potential for both intramuscular adipogenesis and fibrogenesis, leading to the elevation of both marbling and connective tissue content in the resulting meat product. Furthermore, given the bipotent developmental potential of progenitor cells, enhancing their conversion to adipogenesis reduces fibrogenesis, which likely results in the overall improvement of marbling (more intramuscular adipocytes) and tenderness (less connective tissue) of meat. Fibrogenesis is mainly regulated by the transforming growth factor (TGF) β signaling pathway and its regulatory cascade. In addition, extracellular matrix, a part of the intramuscular connective tissue, provides a niche environment for regulating myogenic differentiation of satellite cells and muscle growth. Despite rapid progress, many questions remain in the role of extracellular matrix on muscle development, and factors determining the early differentiation of myogenic, adipogenic and fibrogenic cells, which warrant further studies.

Review Article
© The Animal Consortium 2015 


Intramuscular connective tissue contributes to the background toughness of meat, which is mainly synthesized by intramuscular fibroblasts. Recent studies show that adipocytes and fibroblasts are derived from a common pool of mesenchymal progenitor cells during the early embryonic development. Due to the bipotent developmental potential of these progenitor cells, enhancing their conversion to adipogenesis reduces fibrogenesis, which provides an opportunity to improve marbling and tenderness of meat, thus the overall palatability.


Meat quality is determined by flavor, tenderness, juiciness, color, nutritional value and others. Tender meat, which contains more intramuscular fat and less connective tissue is demanded by consumers. Meat tenderness is determined by both the myofibrillar effects and the presence and cross-linking of connective tissue. Myofibrillar contribution to toughness can be partially addressed by aging carcasses, which results in the fragmentation of myofibrils primarily due to proteolysis by calpains (Koohmaraie and Geesink, Reference Koohmaraie and Geesink2006). On the other hand, postmortem aging is ineffective in improving the tenderness of a meat with high collagen content, due to the resistance of collagen to proteolysis. Thus, meat toughness due to connective tissue is called the ‘background toughness’ of meat (Nishimura, Reference Nishimura2010). Consistently, the longissimus muscle in beef cattle contains low collagen and is tenderer while beef from limb muscles possesses higher collagen content and is tougher (McCormick, Reference McCormick1999; Dubost et al., Reference Dubost, Micol, Meunier, Lethias and Listrat2013a). In addition, the cross-linking of collagen has even greater influence on meat toughness (McCormick, Reference McCormick1994). Because during cooking, collagen is gelatinized, which is hampered due to the presence of cross-linking, contributing to the toughness of meat from old animals (Dubost et al., Reference Dubost, Micol, Picard, Lethias, Andueza, Bauchart and Listrat2013b). The detailed effects of connective tissue structure, collagen cross-linking, and their impacts on meat tenderness have been previous reviewed (Purslow, Reference Purslow, Archile-Contreras and Cha2014).

Intramuscular connective tissue is mainly derived from fibroblasts, which are generated through fibrogenesis, a process referring to the generation of fibroblasts and their synthesis of proteins and other components composing the connective tissue. Fibrogenesis is active during the whole life of animals, particularly during the early developmental stage in utero; connective tissues synthesized inside fetal muscle form primordial perimysium and epimysium of muscle bundles at late gestation (Du et al., Reference Du, Yan, Tong, Zhao and Zhu2010). In humans, fibrosis refers to a state of excessive deposition of collagen and other extracellular matrix proteins, which is often elicited by a pathological condition and becomes noticeable during the recovery period (Liu and Pravia, Reference Liu and Pravia2010). Lysyl oxidase is a rate limiting enzyme catalyzing cross-linking of collagen fibrils (Borg et al., Reference Borg, Klevay, Gay, Siegel and Bergin1985; Huang et al., Reference Huang, Zhao, Yan, Zhu, Long, McCormick, Ford, Nathanielsz and Du2012b). Available studies demonstrated that the content and cross-linking of collagen are frequently correlated to each other, but the turnover of collagen reduces cross-linking (Archile-Contreras et al., Reference Archile-Contreras, Cha, Mandell, Miller and Purslow2010), a process increasing tenderness (Hill, Reference Hill1967; Archile-Contreras et al., Reference Archile-Contreras, Mandell and Purslow2011; Purslow et al., Reference Purslow2012).

Intramuscular fat is considered part of the intramuscular connective tissue, and intramuscular adipogenesis is inseparable from fibrogenesis due to closely related developmental origins. However, knowledge regarding regulatory mechanisms, or specific and effective manipulations to augment progenitor cell differentiation to a particular lineage, such as adipogenesis, remains poorly defined. The intent of this review is to provide an overview of current knowledge regarding intramuscular collagen deposition and associated marbling development, and discuss possible mechanisms regulating mesenchymal progenitor cell differentiation focusing on fibrogenesis, and their impacts on muscle growth and meat quality.

Intramuscular connective tissue structure

Organization of intramuscular connective tissue

All connective tissues (cartilage, bone, blood and interstitial tissue) possess three common components: cells, fibers and ground substance. Extracellular matrix tissue refers to a major portion of intramuscular connective tissues surrounding muscle fibers and other cells, which is composed of collagen, elastin, fibronectin, proteoglycans, and other ground substance components (Purslow, Reference Purslow, Archile-Contreras and Cha2014). Embedded in extracellular matrix and connective tissue, there are abundant fibroblasts, adipocytes, immune cells, preadipocytes, mesenchymal progenitor cells, and other stromal vascular cells. Connective tissue and associated proteins organize muscle structure, connect muscle fibers to the bone for locomotion, and also mediate muscle growth and development (Sanes, Reference Sanes2003; Jenniskens et al., Reference Jenniskens, Koevoet, de Bart, Weinans, Jahr, Verhaar, DeGroot and van Osch2006). The connective tissues surrounding each muscle fiber, termed endomysium, comprised two layers. The inner layer, termed basal lamina, is a 50 to 100 nm thick layer surrounding the sarcolemma, which connects muscle fibers to extracellular niche environment and regulates myogenesis (Wang et al., Reference Wang, Liu, Tsai, Chen, Chang, Tsai, Leu, Zhen, Chai, Chung, Chua, Yen and Yip2014), and muscle growth (Velleman, Reference Velleman1999). Outside of the endomysium, a thin layer of connective tissue, which integrates into thicker layers between muscle bundles, termed perimysium, and surrounding each muscle, termed epimysium. These connective tissues connect muscle fibers and bundles together, and maintain muscle integrity. Intramuscular adipocytes, blood vessels and nerves are integrated into the connective tissue matrix of the muscle.

Connective tissue structure

Collagen is the major component of connective tissue. There are a number of different types of collagens, which are derived from more than 30 genes (Myllyharju and Kivirikko, Reference Myllyharju and Kivirikko2004; Veit et al., Reference Veit, Kobbe, Keene, Paulsson, Koch and Wagener2006; Soderhall et al., Reference Soderhall, Marenholz, Kerscher, Ruschendorf, Esparza-Gordillo, Worm, Gruber, Mayr, Albrecht, Rohde, Schulz, Wahn, Hubner and Lee2007). However, in muscle, types I and III collagen are dominant (Light et al., Reference Light, Champion, Voyle and Bailey1985). The ratio of type I to III may be altered depending on muscle types, locations and animal ages (Listrat et al., Reference Listrat, Picard and Geay1999). In mature bovine muscles, type I collagen is more abundant in perimysium, but type III collagen levels are enriched in the endomysium (Mayne and Sanderson, Reference Mayne and Sanderson1985). In rats, during aging, the proportion of type I collagen increased, while type III collagen decreased (Kovanen and Suominen, Reference Kovanen and Suominen1989); an increase in type I collagen was also observed in the intramuscular connective tissue of beef cattle at around 6 months of age (Listrat et al., Reference Listrat, Picard and Geay1999). Up to now, most studies about connective tissue in muscle have been focused on types I and III collagens (Sato et al., Reference Sato, Ando, Kubota, Origasa, Kawase, Toyohara, Sakaguchi, Nakagawa, Makinodan, Ohtsuki and Kawabata1994; Sato et al., Reference Sato, Sakuma, Ohtsuki and Kawabata1997; Duarte et al., Reference Duarte, Paulino, Das, Wei, Serao, Fu, Harris, Dodson and Du2013).

Each collagen molecule contains three helical polypeptide chains, which are interwined. At both ends, however, non-helical regions termed telopeptide regions are found. Lysyl oxidase is a critical enzyme regulating collagen cross-linking (Siegel and Fu, Reference Siegel and Fu1976; Siegel et al., Reference Siegel, Fu and Chang1976). Lysyl oxidase oxidizes lysine or hydroxylysine in the non-helical portions of collagen molecules to aldehydes, which then react with neighboring collagen molecules to form divalent bonds. Therefore, the presence of lysine and hydroxylysine in the non-helical regions is critical in determining cross-linking development (Robins, Reference Robins2007). The degree of collagen cross-linking differs in animals of different breeds. In our study with Wagyu and Angus cattle, we found that the collagen content and cross-linking are higher in Wagyu, which correlates with less soluble collagen content (Duarte et al., Reference Duarte, Paulino, Das, Wei, Serao, Fu, Harris, Dodson and Du2013). We also observed that early nutrition affects collagen content and cross-linking in sheep (Huang et al., Reference Huang, Yan, Zhu, McCormick, Ford, Nathanielsz and Du2010). In addition, collagens of different muscle types have various degrees of cross-linking, with the collagen in longissimus muscle having less cross-linking than biceps muscle (Dubost et al., Reference Dubost, Micol, Meunier, Lethias and Listrat2013a), correlated with meat tenderness. Collagen cross-linking is a slow process, which increases as animals age, and the high degree of cross-linking is one of the primary reasons for the toughness of meat from old animals. On the other hand, collagens undergo consistent turnover, albeit slower than other proteins. Because newly synthesized collagens do not contain cross-linking, factors that enhance collagen turnover, reduce cross-linking and improve meat tenderness (Purslow, Reference Purslow, Archile-Contreras and Cha2014). Indeed, cross-linking was reduced and soluble collagen content was raised in compensatory growing pigs (Kristensen et al., Reference Kristensen, Therkildsen, Riis, Sørensen, Oksbjerg, Purslow and Ertbjerg2002). Collagen turnover, or remodeling, is regulated by metalloproteinases (Woessner, Reference Woessner1991; Murphy, Reference Murphy2010). The expression of metalloproteinases and their inhibitors, the tissue inhibitors of metalloproteinases, are regulated by a number of factors (Clark et al., Reference Clark, Swingler, Sampieri and Edwards2008), such as inflammation and oxidative stress, which affect cross-linking and meat tenderness (Purslow, Reference Purslow, Archile-Contreras and Cha2014).

Development of connective tissue

Fibrogenic cells and adipocytes share common progenitor cells

During early skeletal muscle development, mesenchymal stem cells first diverge to either myogenic or non-myogenic lineages. Myogenic progenitors further develop into muscle fibers and satellite cells, whereas non-myogenic progenitor cells develop into the stromal-vascular fraction of mature skeletal muscle in which resides adipocytes, fibroblasts and resident mesenchymal progenitor cells (Du et al., Reference Du, Huang, Das, Yang, Duarte, Dodson and Zhu2013). These non-myogenic progenitors have adipogenic and fibrogenic capacity, as well as osteogenic and chondrogenic potential (Joe et al., Reference Joe, Yi, Natarajan, Le Grand, So, Wang, Rudnicki and Rossi2010; Wosczyna et al., Reference Wosczyna, Biswas, Cogswell and Goldhamer2012). These cells are mainly located in the stromal-vascular fraction of skeletal muscle and are distinct from satellite cells (Joe et al., Reference Joe, Yi, Natarajan, Le Grand, So, Wang, Rudnicki and Rossi2010; Uezumi et al., Reference Uezumi, Ikemoto-Uezumi and Tsuchida2010). Platelet-derived growth factor receptor α (PDGFRα) is a reliable marker for separating these cells, and Sca-1+CD34+ appears to label the same cell population (Joe et al., Reference Joe, Yi, Natarajan, Le Grand, So, Wang, Rudnicki and Rossi2010; Uezumi et al., Reference Uezumi, Ikemoto-Uezumi and Tsuchida2010, Reference Uezumi, Ito, Morikawa, Shimizu, Yoneda, Segawa, Yamaguchi, Ogawa, Matev, Miyagoe-Suzuki, Takeda, Tsujikawa, Tsuchida, Yamamoto and Fukada2011 and Reference Uezumi, Fukada, Yamamoto, Takeda and Tsuchida2014).

The notion that mesenchymal progenitor cells as the common sources of adipogenic and fibrogenic cells are further proven by the co-expression of PDGFRα with fibrogenic markers (Murphy et al., Reference Murphy, Lawson, Mathew, Hutcheson and Kardon2011), or PDGFRα with adipogenic markers (Yang et al., Reference Yang, Liang, Rogers, Zhao, Zhu and Du2013). Transcription factor 4 (TCF4), also known as transcription factor 7-like 2 (Tcf7l2), was first found to be related with limb development by interacting with Wnt signaling pathway (Cho and Dressler, Reference Cho and Dressler1998). Subsequent studies demonstrate TCF4 as a fibrogenic marker (Kardon et al., Reference Kardon, Harfe and Tabin2003; Mathew et al., Reference Mathew, Hansen, Merrell, Murphy, Lawson, Hutcheson, Hansen, Angus-Hill and Kardon2011). A portion of TCF4+ fibroblasts also express PDGFRα (Murphy et al., Reference Murphy, Lawson, Mathew, Hutcheson and Kardon2011), showing the intrinsic relationship between mesenchymal progenitor cells and TCF4+ fibroblasts. Similarly, in our previous studies, we detected the co-expression of PDGFRα with ZFP423, a marker of adipogenic commitment (Yang et al., Reference Yang, Liang, Rogers, Zhao, Zhu and Du2013). The lack of TCF4+ and ZFP423 co-expressed cells show the divergence of the fibrogenic and adipogenic lineages during progenitor differentiation.

Mechanisms regulating fibrogenesis

Transforming growth factor (TGF)-β is the most important profibrogenic cytokine (Liu and Pravia, Reference Liu and Pravia2010). TGF superfamily contains several structurally related subfamilies, including TGF-β, bone morphogenetic proteins and activin. Three isoforms of TGF-β have been identified, which are TGF-β1, TGF-β2 and TGF-β3. The TGF-β1 isoform is primarily expressed in endothelial cells, fibroblasts, hematopoietic cells and smooth muscle cells; TGF-β2 mainly exists in epithelial cells and neurons; and TGF-β3 is specifically expressed in mesenchymal cells (Ghosh et al., Reference Ghosh, Murphy, Turner, Khwaja, Halka, Kielty and Walker2005). All TGF-β isoforms activate down-stream SMAD signaling (Attisano and Wrana, Reference Attisano and Wrana1996; Letterio and Roberts, Reference Letterio and Roberts1998). The SMAD family contains five receptor-regulated SMAD (R-SMAD 1, 2, 3, 5 and 8), a common SMAD (Co-SMAD 4), and two inhibitor SMAD (I-SMAD 6 and 7) (Moustakas et al., Reference Moustakas, Souchelnytskyi and Heldin2001). The ligand, TGF-β, first binds to TGF-β receptor II (TβRII), which then recruits and activates TβRI. Then SMAD2 and SMAD3 are phosphorylated and subsequently bind to SMAD4 (Suwanabol et al., Reference Suwanabol, Kent and Liu2011), and the resulting SMAD complex is translocated into the nucleus where it binds to SMAD-specific binding elements of target genes, thereby activating the expression of fibrogenic genes including procollagen and enzymes catalyzing collagen cross-linking (Massague and Chen, Reference Massague and Chen2000). As an anti-inflammatory cytokine, TGF-β signaling is enhanced by inflammation (Bhatnagar et al., Reference Bhatnagar, Panguluri, Gupta, Dahiya, Lundy and Kumar2010; Voloshenyuk et al., Reference Voloshenyuk, Hart, Khoutorova and Gardner2011), while inhibited by anti-inflammatory factors (Wang et al., Reference Wang, Dumont and Rudnicki2012).

Connective tissue growth factor (CTGF) is a crucial switch to regulate downstream fibrotic progress (Grotendorst, Reference Grotendorst1997; Leask et al., Reference Leask, Denton and Abraham2004). CTGF is a member of CCN family, which are cysteine rich proteins. CTGF gene expression is induced by TGF-β-activated Smad3 binding to its promoter region (Denton and Abraham, Reference Denton and Abraham2001; Holmes et al., Reference Holmes, Abraham, Sa, Shiwen, Black and Leask2001). Then, CTGF directly stimulates fibroblast proliferation and ECM deposition (Shi-Wen et al., Reference Shi-Wen, Leask and Abraham2008; Morales et al., Reference Morales, Cabello-Verrugio, Santander, Cabrera, Goldschmeding and Brandan2011). Wingless/int (Wnt) signaling pathway plays a crucial role in cell fate commitment (Dorsky et al., Reference Dorsky, Moon and Raible1998; Ross et al., Reference Ross, Hemati, Longo, Bennett, Lucas, Erickson and MacDougald2000), and synergizes with TGF-β signaling to promote connective tissue synthesis and fibrosis (Brack et al., Reference Brack, Conboy, Roy, Lee, Kuo, Keller and Rando2007; Zhou et al., Reference Zhou, Liu, Kahn, Ann, Han, Wang, Nguyen, Flodby, Zhong, Krishnaveni, Liebler, Minoo, Crandall and Borok2012; Cisternas et al., Reference Cisternas, Vio and Inestrosa2014).

Ski/sno family includes ski and sno, which has four distinct isoforms SnoN, SnoN2, SnoA and Snol (Nomura et al., Reference Nomura, Sasamoto, Ishii, Date, Matsui and Ishizaki1989; Pearson-White, Reference Pearson-White1993; Pelzer et al., Reference Pelzer, Lyons, Kim and Moreadith1996). Ski/sno family acts as negative regulators of TGF-β1 pathway by functioning on the downstream signal molecules R-smad/Co-smad complex (Luo, Reference Luo2004; Deheuninck and Luo, Reference Deheuninck and Luo2009; Jahchan and Luo, Reference Jahchan and Luo2010), thus reducing connective tissue deposition.

MicroRNAs regulate cell differentiation through inhibiting the expression of target genes. MiR-101a inhibits fibrosis by targeting the TβRI on cardiac fibroblasts (Zhao et al., Reference Zhao, Wang, Liao, Zeng, Li, Hu, Liu, Meng, Qian, Zhang, Guan, Feng, Zhou, Du and Chen2015). High glucose increases the activity of transcriptional co-activator p300, which subsequently enhances the activity of TGFβ pathway by inducing Smad2 acetylation (Bugyei-Twum et al., Reference Bugyei-Twum, Advani, Advani, Zhang, Thai, Kelly and Connelly2014). Besides, ERK5, one of the MAPK family members, is a critical regulator in TGF-β1-induced lung fibrosis by enhancing Smad3 acetylation (Kim et al., Reference Kim, Lim and Woo2013). A number of cytokines and growth factors, which are involved in the regulation of fibrogenesis are listed in Table 1.

Table 1 Factors enhancing and decreasing intramuscular fibrogenesis

CTGF=connective tissue growth factor; FGF-2=basic fibroblast growth factor; MMPs=matrix metablloproteinase; PDGF=platelet-derived growth factor; TGFβ=tumor growth factor β; TIMP=tissue inhibitor of metalloproteinase; Wnts=wingless and ints.

Antagonistic effects of adipogenesis on fibrogenesis

Because fibrogenesis and adipogenesis are considered as a competitive process, enhancing adipogenesis reduces fibrogenesis. Adipogenesis can be separated into two steps, the commitment of progenitors to preadipocytes, and the differentiation of preadipocytes to mature adipocytes. Quite recently, Zfp423 was identified as the key regulator committing progenitors to preadipocytes; in addition, Zfp423 promotes the expression of peroxisome proliferator-activated receptor γ, the crucial transcription factor inducing the conversion of preadipocytes to adipocytes (Gupta et al., Reference Gupta, Arany, Seale, Mepani, Ye, Conroe, Roby, Kulaga, Reed and Spiegelman2010; Gupta et al., Reference Gupta, Mepani, Kleiner, Lo, Khandekar, Cohen, Frontini, Bhowmick, Ye, Cinti and Spiegelman2012). Importantly, in cattle mesenchymal progenitor cells, the expression of Zfp423 is negatively correlated with TGF-β1 expression, indicating the mutual exclusion of adipogenesis and fibrogenesis (Huang et al., Reference Huang, Das, Yang, Zhu and Du2012a).

Connective tissue and muscle development

Satellite cells are critical for muscle growth and regeneration. They are wedged between the basal lamina and the plasma membrane (sarcolemma) of skeletal muscle fibers. Extracellular matrix together with growth factors and cytokines sequestered inside and those secreted by interstitial cells, forms the niche environment needed for satellite cell quiescence, activation, migration, myogenic differentiation and muscle development (Rhoads et al., Reference Rhoads, Fernyhough, Liu, McFarland, Velleman, Hausman and Dodson2009; Dodson et al., Reference Dodson, Hausman, Guan, Du, Rasmussen, Poulos, Mir, Bergen, Fernyhough, McFarland, Rhoads, Soret, Reecy, Velleman and Jiang2010; Murphy et al., Reference Murphy, Lawson, Mathew, Hutcheson and Kardon2011; Urciuolo et al., Reference Urciuolo, Quarta, Morbidoni, Gattazzo, Molon, Grumati, Montemurro, Tedesco, Blaauw, Cossu, Vozzi, Rando and Bonaldo2013).

Muscle regeneration involves extensive proliferation and myogenic differentiation of satellite cells. Shortly after muscle injury, both satellite cells and non-myogenic progenitor cells are activated and proliferate; non-myogenic progenitor cells stimulate satellite cell proliferation and facilitate muscle regeneration (Joe et al., Reference Joe, Yi, Natarajan, Le Grand, So, Wang, Rudnicki and Rossi2010; Murphy et al., Reference Murphy, Lawson, Mathew, Hutcheson and Kardon2011). In addition, intramuscular fibroblasts particularly promote slow myogenesis, thus affecting muscle fiber type composition and overall maturation during muscle development (Mathew et al., Reference Mathew, Hansen, Merrell, Murphy, Lawson, Hutcheson, Hansen, Angus-Hill and Kardon2011). Extracellular component, collagen VI, regulates satellite cell self-renewal and differentiation (Urciuolo et al., Reference Urciuolo, Quarta, Morbidoni, Gattazzo, Molon, Grumati, Montemurro, Tedesco, Blaauw, Cossu, Vozzi, Rando and Bonaldo2013). Besides, other components of extracellular matrix, such as proteoglycan, regulate proliferation and differentiation of satellite cells (Zhang et al., Reference Zhang, Nestor, McFarland and Velleman2007). Decorin, a small leucine-rich proteoglycan, traps TGFβ to regulate satellite cell activation and muscle growth (Li et al., Reference Li, McFarland and Velleman2006 and Reference Li, McFarland and Velleman2008).

Extracellular matrix also interacts with a number of growth factors, including TGFβ, hepatocyte growth factor, fibroblast growth factor 2, myostatin and others to either promote or inhibit muscle growth (Yamaguchi et al., Reference Yamaguchi, Mann and Ruoslahti1990; Rapraeger et al., Reference Rapraeger, Krufka and Olwin1991; Allen et al., Reference Allen, Sheehan, Taylor, Kendall and Rice1995; Miura et al., Reference Miura, Kishioka, Wakamatsu, Hattori, Hennebry, Berry, Sharma, Kambadur and Nishimura2006; Kishioka et al., Reference Kishioka, Thomas, Wakamatsu, Hattori, Sharma, Kambadur and Nishimura2008). Table 2 lists selected growth factors known to interact with extracellular matrix and regulate muscle growth.

Table 2 Growth factors associated with extracellular matrix and associated cells to regulate activation of satellite cells

EGF=epithelial growth factor; FGF-2=fibroblast growth factor-2; HGF/SF=hepatocyte growth factor/scatter factor; IGF=insulin growth factor; PDGF-BB=platelet-derived growth factor-BB; SDF-1=stromal-derived factor-1; TGFβ=transforming growth factor β.


Intramuscular connective tissue regulates muscle growth and development, and also is the site for intramuscular fat (marbling) deposition. The abundance and cross-linking of intramuscular connective tissue contribute to the background toughness of meat. Connective tissue is mainly synthesized by intramuscular fibroblasts. Non-myogenic mesenchymal progenitor cells are the common source of fibroblasts and adipocytes. Strengthening progenitor cell formation and proliferation enhances both intramuscular adipogenesis and fibrogenesis, while enhancing progenitor differentiation to adipogenesis reduces fibrogenesis, resulting in the overall improvement of marbling and tenderness of meat. Fibrogenesis is mainly regulated by the TGFβ signaling pathway, and a number of factors affect connective tissue deposition via altering TGFβ signaling. Extracellular matrix, a part of the intramuscular connective tissue, provides a niche environment to regulate myogenic differentiation of satellite cells and muscle growth. Despite rapid progress in our understanding of mechanisms regulating fibrogenesis, many questions remain on the synthesis of intramuscular connective tissue and the role of extracellular matrix in muscle development, which warrants further studies.


This project was supported by Agriculture and Food Research Initiative Competitive Grant No. 2015-67015-23219 from the USDA National Institute of Food and Agriculture, and NIH R01 HD067449.


Allen, RE, Sheehan, SM, Taylor, RG, Kendall, TL and Rice, GM 1995. Hepatocyte growth factor activates quiescent skeletal muscle satellite cells in vitro. Journal of Cellular Physiology 165, 307312.CrossRefGoogle ScholarPubMed
Archile-Contreras, AC, Cha, MC, Mandell, IB, Miller, SP and Purslow, PP 2011. Vitamins E and C may increase collagen turnover by intramuscular fibroblasts. Potential for improved meat quality. Journal of Agricultural and Food Chemistry 59, 608614.CrossRefGoogle Scholar
Archile-Contreras, AC, Mandell, IB and Purslow, PP 2010. Disparity of dietary effects on collagen characteristics and toughness between two beef muscles. Meat Science 86, 491497.CrossRefGoogle ScholarPubMed
Attisano, L and Wrana, JL 1996. Signal transduction by members of the transforming growth factor-beta superfamily. Cytokine & Growth Factor Reviews 7, 327339.CrossRefGoogle ScholarPubMed
Balcerzak, D, Querengesser, L, Dixon, WT and Baracos, VE 2001. Coordinate expression of matrix-degrading proteinases and their activators and inhibitors in bovine skeletal muscle. Journal of Animal Science 79, 94107.CrossRefGoogle ScholarPubMed
Bhatnagar, S, Panguluri, SK, Gupta, SK, Dahiya, S, Lundy, RF and Kumar, A 2010. Tumor necrosis factor-alpha regulates distinct molecular pathways and gene networks in cultured skeletal muscle cells. PLoS One 5, e13262.CrossRefGoogle ScholarPubMed
Borg, TK, Klevay, LM, Gay, RE, Siegel, R and Bergin, ME 1985. Alteration of the connective tissue network of striated muscle in copper deficient rats. Journal of Molecular and Cellular Cardiology 17, 11731183.CrossRefGoogle ScholarPubMed
Brack, AS, Conboy, MJ, Roy, S, Lee, M, Kuo, CJ, Keller, C and Rando, TA 2007. Increased Wnt signaling during aging alters muscle stem cell fate and increases fibrosis. Science 317, 807810.CrossRefGoogle ScholarPubMed
Brzoska, E, Kowalewska, M, Markowska-Zagrajek, A, Kowalski, K, Archacka, K, Zimowska, M, Grabowska, I, Czerwinska, AM, Czarnecka-Gora, M, Streminska, W, Janczyk-Ilach, K and Ciemerych, MA 2012. Sdf-1 (CXCL12) improves skeletal muscle regeneration via the mobilisation of Cxcr4 and CD34 expressing cells. Biology of the Cell 104, 722737.CrossRefGoogle ScholarPubMed
Bugyei-Twum, A, Advani, A, Advani, SL, Zhang, Y, Thai, K, Kelly, DJ and Connelly, KA 2014. High glucose induces Smad activation via the transcriptional coregulator p300 and contributes to cardiac fibrosis and hypertrophy. Cardiovascular Diabetology 13, 89.CrossRefGoogle ScholarPubMed
Cho, EA and Dressler, GR 1998. TCF-4 binds beta-catenin and is expressed in distinct regions of the embryonic brain and limbs. Mechanisms of Development 77, 918.CrossRefGoogle ScholarPubMed
Cisternas, P, Vio, CP and Inestrosa, NC 2014. Role of Wnt signaling in tissue fibrosis, lessons from skeletal muscle and kidney. Current Molecular Medicine 14, 510522.CrossRefGoogle ScholarPubMed
Clark, IM, Swingler, TE, Sampieri, CL and Edwards, DR 2008. The regulation of matrix metalloproteinases and their inhibitors. The International Journal of Biochemistry & Cell Biology 40, 13621378.CrossRefGoogle ScholarPubMed
Deheuninck, J and Luo, K 2009. Ski and SnoN, potent negative regulators of TGF-beta signaling. Cell Research 19, 4757.CrossRefGoogle ScholarPubMed
Denton, CP and Abraham, DJ 2001. Transforming growth factor-beta and connective tissue growth factor: key cytokines in scleroderma pathogenesis. Current Opinion in Rheumatology 13, 505511.CrossRefGoogle ScholarPubMed
Dodson, MV, Hausman, GJ, Guan, L, Du, M, Rasmussen, TP, Poulos, SP, Mir, P, Bergen, WG, Fernyhough, ME, McFarland, DC, Rhoads, RP, Soret, B, Reecy, JM, Velleman, SG and Jiang, Z 2010. Skeletal muscle stem cells from animals I. Basic cell biology. International Journal of Biological Science 6, 465474.CrossRefGoogle ScholarPubMed
Dorsky, RI, Moon, RT and Raible, DW 1998. Control of neural crest cell fate by the Wnt signalling pathway. Nature 396, 370373.CrossRefGoogle ScholarPubMed
Doumit, ME, Cook, DR and Merkel, RA 1993. Fibroblast growth factor, epidermal growth factor, insulin-like growth factors, and platelet-derived growth factor-BB stimulate proliferation of clonally derived porcine myogenic satellite cells. Journal of Cell Physiology 157, 326332.CrossRefGoogle ScholarPubMed
Du, M, Huang, Y, Das, AK, Yang, Q, Duarte, MS, Dodson, MV and Zhu, M-J 2013. Meat Science And Muscle Biology Symposium: manipulating mesenchymal progenitor cell differentiation to optimize performance and carcass value of beef cattle. Journal of Animal Science 91, 14191427.CrossRefGoogle ScholarPubMed
Du, M, Yan, X, Tong, JF, Zhao, JX and Zhu, MJ 2010. Maternal obesity, inflammation, and fetal skeletal muscle development. Biology of Reproduction 82, 412.CrossRefGoogle ScholarPubMed
Duarte, MS, Paulino, PV, Das, AK, Wei, S, Serao, NV, Fu, X, Harris, SM, Dodson, MV and Du, M 2013. Enhancement of adipogenesis and fibrogenesis in skeletal muscle of Wagyu compared with Angus cattle. Journal of Animal Science 91, 29382946.CrossRefGoogle ScholarPubMed
Dubost, A, Micol, D, Meunier, B, Lethias, C and Listrat, A 2013a. Relationships between structural characteristics of bovine intramuscular connective tissue assessed by image analysis and collagen and proteoglycan content. Meat Science 93, 378386.CrossRefGoogle ScholarPubMed
Dubost, A, Micol, D, Picard, B, Lethias, C, Andueza, D, Bauchart, D and Listrat, A 2013b. Structural and biochemical characteristics of bovine intramuscular connective tissue and beef quality. Meat Science 95, 555561.CrossRefGoogle ScholarPubMed
Ghosh, J, Murphy, MO, Turner, N, Khwaja, N, Halka, A, Kielty, CM and Walker, MG 2005. The role of transforming growth factor beta1 in the vascular system. Cardiovascular Pathology 14, 2836.CrossRefGoogle ScholarPubMed
Grotendorst, GR 1997. Connective tissue growth factor: a mediator of TGF-beta action on fibroblasts. Cytokine & Growth Factor Reviews 8, 171179.CrossRefGoogle ScholarPubMed
Gupta, RK, Arany, Z, Seale, P, Mepani, RJ, Ye, L, Conroe, HM, Roby, YA, Kulaga, H, Reed, RR and Spiegelman, BM 2010. Transcriptional control of preadipocyte determination by Zfp423. Nature 464, 619623.CrossRefGoogle ScholarPubMed
Gupta, RK, Mepani, RJ, Kleiner, S, Lo, JC, Khandekar, MJ, Cohen, P, Frontini, A, Bhowmick, DC, Ye, L, Cinti, S and Spiegelman, BM 2012. Zfp423 expression identifies committed preadipocytes and localizes to adipose endothelial and perivascular cells. Cell Metabolism 15, 230239.CrossRefGoogle ScholarPubMed
Haugk, KL, Roeder, RA, Garber, MJ and Schelling, GT 1995. Regulation of muscle cell proliferation by extracts from crushed muscle. Journal of Animal Science 73, 19721981.CrossRefGoogle ScholarPubMed
Hill, F 1967. The chemical composition of muscles from steers which experienced compensatory growth. Journal of the Science of Food Agriculture 18, 164166.CrossRefGoogle ScholarPubMed
Holmes, A, Abraham, DJ, Sa, S, Shiwen, X, Black, CM and Leask, A 2001. CTGF and SMADs, maintenance of scleroderma phenotype is independent of SMAD signaling. Journal of Biological Chemistry 276, 1059410601.CrossRefGoogle ScholarPubMed
Huang, Y, Das, AK, Yang, QY, Zhu, MJ and Du, M 2012a. Zfp423 promotes adipogenic differentiation of bovine stromal vascular cells. PLoS One 7, e47496.CrossRefGoogle ScholarPubMed
Huang, Y, Yan, X, Zhu, MJ, McCormick, RJ, Ford, SP, Nathanielsz, PW and Du, M 2010. Enhanced transforming growth factor-beta signaling and fibrogenesis in ovine fetal skeletal muscle of obese dams at late gestation. American Journal of Physiology – Endocrinology and Metabolism 298, E1254E1260.CrossRefGoogle ScholarPubMed
Huang, Y, Zhao, JX, Yan, X, Zhu, MJ, Long, NM, McCormick, RJ, Ford, SP, Nathanielsz, PW and Du, M 2012b. Maternal obesity enhances collagen accumulation and cross-linking in skeletal muscle of ovine offspring. PLoS One 7, e31691.CrossRefGoogle ScholarPubMed
Iannaccone, S, Quattrini, A, Smirne, S, Sessa, M, de Rino, F, Ferini-Strambi, L and Nemni, R 1995. Connective tissue proliferation and growth factors in animal models of Duchenne muscular dystrophy. Journal of the Neurological Sciences 128, 3644.CrossRefGoogle ScholarPubMed
Jahchan, NS and Luo, K 2010. SnoN in mammalian development, function and diseases. Current Opinion in Pharmacology 10, 670675.CrossRefGoogle ScholarPubMed
Jenniskens, YM, Koevoet, W, de Bart, ACW, Weinans, H, Jahr, H, Verhaar, JAN, DeGroot, J and van Osch, GJVM 2006. Biochemical and functional modulation of the cartilage collagen network by IGF1, TGF beta 2 and FGF2. Osteoarthritis and Cartilage 14, 11361146.CrossRefGoogle ScholarPubMed
Joe, AW, Yi, L, Natarajan, A, Le Grand, F, So, L, Wang, J, Rudnicki, MA and Rossi, FM 2010. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nature Cell Biology 12, 153163.CrossRefGoogle ScholarPubMed
Kardon, G, Harfe, BD and Tabin, CJ 2003. A Tcf4-positive mesodermal population provides a prepattern for vertebrate limb muscle patterning. Developmental Cell 5, 937944.CrossRefGoogle ScholarPubMed
Kim, S, Lim, JH and Woo, CH 2013. ERK5 inhibition ameliorates pulmonary fibrosis via regulating Smad3 acetylation. American Journal of Pathology 183, 17581768.CrossRefGoogle ScholarPubMed
Kishioka, Y, Thomas, M, Wakamatsu, Ji, Hattori, A, Sharma, M, Kambadur, R and Nishimura, T 2008. Decorin enhances the proliferation and differentiation of myogenic cells through suppressing myostatin activity. Journal of Cellular Physiology 215, 856867.CrossRefGoogle ScholarPubMed
Koohmaraie, M and Geesink, GH 2006. Contribution of postmortem muscle biochemistry to the delivery of consistent meat quality with particular focus on the calpain system. Meat Science 74, 3443.CrossRefGoogle ScholarPubMed
Kovanen, V and Suominen, H 1989. Age- and training-related changes in the collagen metabolism of rat skeletal muscle. European Journal of Applied Physiology and Occupational Physiology 58, 765771.CrossRefGoogle ScholarPubMed
Kristensen, L, Therkildsen, M, Riis, B, Sørensen, MT, Oksbjerg, N, Purslow, P and Ertbjerg, P 2002. Dietary-induced changes of muscle growth rate in pigs: effects on in vivo and postmortem muscle proteolysis and meat quality. Journal of Animal Science 80, 28622871.CrossRefGoogle ScholarPubMed
Leask, A, Denton, CP and Abraham, DJ 2004. Insights into the molecular mechanism of chronic fibrosis: the role of connective tissue growth factor in scleroderma. Journal of Investigative Dermatology 122, 16.CrossRefGoogle ScholarPubMed
Letterio, JJ and Roberts, AB 1998. Regulation of immune responses by TGF-beta. Annual Review Immunology 16, 137161.CrossRefGoogle ScholarPubMed
Li, X, McFarland, DC and Velleman, SG 2006. Effect of transforming growth factor-beta on decorin and beta1 integrin expression during muscle development in chickens. Poultry Science 85, 326332.CrossRefGoogle ScholarPubMed
Li, X, McFarland, DC and Velleman, SG 2008. Extracellular matrix proteoglycan decorin-mediated myogenic satellite cell responsiveness to transforming growth factor-beta1 during cell proliferation and differentiation Decorin and transforming growth factor-beta1 in satellite cells. Domestic Animal Endocrinology 35, 263273.CrossRefGoogle ScholarPubMed
Light, N, Champion, AE, Voyle, C and Bailey, AJ 1985. The role of epimysial, perimysial and endomysial collagen in determining texture in six bovine muscles. Meat Science 13, 137149.CrossRefGoogle ScholarPubMed
Listrat, A, Picard, B and Geay, Y 1999. Age-related changes and location of type I, III, IV, V and VI collagens during development of four foetal skeletal muscles of double-muscled and normal bovine animals. Tissue Cell 31, 1727.CrossRefGoogle ScholarPubMed
Liu, RM and Pravia, KAG 2010. Oxidative stress and glutathione in TGF-beta-mediated fibrogenesis. Free Radical Biology and Medicine 48, 115.CrossRefGoogle ScholarPubMed
Liu, X, Sun, Y, Weinberg, RA and Lodish, HF 2001. Ski/Sno and TGF-beta signaling. Cytokine & Growth Factor Reviews 12, 18.CrossRefGoogle ScholarPubMed
Luo, K 2004. Ski and SnoN: negative regulators of TGF-beta signaling. Current Opinion in Genetics & Development 14, 6570.CrossRefGoogle ScholarPubMed
Makihara, N, Arimura, K, Ago, T, Tachibana, M, Nishimura, A, Nakamura, K, Matsuo, R, Wakisaka, Y, Kuroda, J, Sugimori, H, Kamouchi, M and Kitazono, T 2015. Involvement of platelet-derived growth factor receptor beta in fibrosis through extracellular matrix protein production after ischemic stroke. Experimental Neurology 264, 127134.CrossRefGoogle ScholarPubMed
Massague, J and Chen, YG 2000. Controlling TGF-beta signaling. Genes & Development 14, 627644.Google ScholarPubMed
Mathew, SJ, Hansen, JM, Merrell, AJ, Murphy, MM, Lawson, JA, Hutcheson, DA, Hansen, MS, Angus-Hill, M and Kardon, G 2011. Connective tissue fibroblasts and Tcf4 regulate myogenesis. Development 138, 371384.CrossRefGoogle ScholarPubMed
Mau, M, Kalbe, C, Wollenhaupt, K, Nurnberg, G and Rehfeldt, C 2008. IGF-I- and EGF-dependent DNA synthesis of porcine myoblasts is influenced by the dietary isoflavones genistein and daidzein. Domestic Animal Endocrinology 35, 281289.CrossRefGoogle ScholarPubMed
Mayne, R and Sanderson, RD 1985. The extracellular matrix of skeletal muscle. Collagen and Related Research 5, 449468.CrossRefGoogle ScholarPubMed
McCormick, RJ 1994. The flexibility of the collagen compartment of muscle. Meat Science 36, 7991.CrossRefGoogle ScholarPubMed
McCormick, RJ 1999. Extracellular modifications to muscle collagen: implications for meat quality. Poultry Science 78, 785791.CrossRefGoogle ScholarPubMed
McFarland, DC, Pesall, JE and Gilkerson, KK 1993. The influence of growth factors on turkey embryonic myoblasts and satellite cells in vitro. General and Comparative Endocrinology 89, 415424.CrossRefGoogle ScholarPubMed
Miura, T, Kishioka, Y, Wakamatsu, J-i, Hattori, A, Hennebry, A, Berry, CJ, Sharma, M, Kambadur, R and Nishimura, T 2006. Decorin binds myostatin and modulates its activity to muscle cells. Biochemical and Biophysical Research Communications 340, 675680.CrossRefGoogle ScholarPubMed
Morales, MG, Cabello-Verrugio, C, Santander, C, Cabrera, D, Goldschmeding, R and Brandan, E 2011. CTGF/CCN-2 over-expression can directly induce features of skeletal muscle dystrophy. Journal of Pathology 225, 490501.CrossRefGoogle ScholarPubMed
Moustakas, A, Souchelnytskyi, S and Heldin, CH 2001. Smad regulation in TGF-beta signal transduction. Journal of Cell Science 114, 43594369.Google ScholarPubMed
Murphy, G 2010. Fell‐Muir Lecture: metalloproteinases: from demolition squad to master regulators. International Journal of Experimental Pathology 91, 303313.CrossRefGoogle ScholarPubMed
Murphy, MM, Lawson, JA, Mathew, SJ, Hutcheson, DA and Kardon, G 2011. Satellite cells, connective tissue fibroblasts and their interactions are crucial for muscle regeneration. Development 138, 36253637.CrossRefGoogle ScholarPubMed
Myllyharju, J and Kivirikko, KI 2004. Collagens, modifying enzymes and their mutations in humans, flies and worms. Trends in Genetics 20, 3343.CrossRefGoogle ScholarPubMed
Nishimura, T 2010. The role of intramuscular connective tissue in meat texture. Animal Science Journal 81, 2127.CrossRefGoogle ScholarPubMed
Nomura, N, Sasamoto, S, Ishii, S, Date, T, Matsui, M and Ishizaki, R 1989. Isolation of human cDNA clones of ski and the ski-related gene, sno. Nucleic Acids Research 17, 54895500.CrossRefGoogle ScholarPubMed
Pearson-White, S 1993. SnoI, a novel alternatively spliced isoform of the ski protooncogene homolog, sno. Nucleic Acids Research 21, 46324638.CrossRefGoogle ScholarPubMed
Pelzer, T, Lyons, GE, Kim, S and Moreadith, RW 1996. Cloning and characterization of the murine homolog of the sno proto-oncogene reveals a novel splice variant. Developmental Dynamics 205, 114125.3.0.CO;2-L>CrossRefGoogle ScholarPubMed
Poncelet, AC and Schnaper, HW 2001. Sp1 and Smad proteins cooperate to mediate transforming growth factor-beta 1-induced alpha 2(I) collagen expression in human glomerular mesangial cells. Journal of Biological Chemistry 276, 69836992.CrossRefGoogle ScholarPubMed
Purslow, PP 2014. New developments on the role of intramuscular connective tissue in meat toughness. Annual Review of Food Science and Technology 5, 133153.CrossRefGoogle ScholarPubMed
Purslow, P, Archile-Contreras, A and Cha, M 2012. Meat Science And Muscle Biology Symposium: manipulating meat tenderness by increasing the turnover of intramuscular connective tissue. Journal of Animal Science 90, 950959.CrossRefGoogle ScholarPubMed
Rapraeger, AC, Krufka, A and Olwin, BB 1991. Requirement of heparan sulfate for bFGF-mediated fibroblast growth and myoblast differentiation. Science 252, 17051708.CrossRefGoogle ScholarPubMed
Rhoads, RP, Fernyhough, ME, Liu, X, McFarland, DC, Velleman, SG, Hausman, GJ and Dodson, MV 2009. Extrinsic regulation of domestic animal-derived myogenic satellite cells II. Domestic Animal Endocrinology 36, 111126.CrossRefGoogle ScholarPubMed
Robins, SP 2007. Biochemistry and functional significance of collagen cross-linking. Biochemical Society Transactions 35, 849852.CrossRefGoogle ScholarPubMed
Roe, JA, Harper, JM and Buttery, PJ 1989. Protein metabolism in ovine primary muscle cultures derived from satellite cells – effects of selected peptide hormones and growth factors. Journal of Endocrinology 122, 565571.CrossRefGoogle ScholarPubMed
Ross, SE, Hemati, N, Longo, KA, Bennett, CN, Lucas, PC, Erickson, RL and MacDougald, OA 2000. Inhibition of adipogenesis by Wnt signaling. Science 289, 950953.CrossRefGoogle ScholarPubMed
Sanes, JR 2003. The basement membrane/basal lamina of skeletal muscle. Journal of Biological Chemistry 278, 1260112604.CrossRefGoogle ScholarPubMed
Sato, K, Ando, M, Kubota, S, Origasa, K, Kawase, H, Toyohara, H, Sakaguchi, M, Nakagawa, T, Makinodan, Y, Ohtsuki, K and Kawabata, M 1997. Involvement of type V collagen in softening of fish muscle during short-term chilled storage. Journal of Agricultural and Food Chemistry 45, 343348.CrossRefGoogle Scholar
Sato, K, Sakuma, A, Ohtsuki, K and Kawabata, M 1994. Subunit composition of Eel (Anguilla-Japonica) type-V collagen – evidence for existence of a novel 4Th Alpha-4(V) chain. Journal of Agricultural and Food Chemistry 42, 675678.CrossRefGoogle Scholar
Sheehan, SM and Allen, RE 1999. Skeletal muscle satellite cell proliferation in response to members of the fibroblast growth factor family and hepatocyte growth factor. Journal of Cell Physiology 181, 499506.3.0.CO;2-1>CrossRefGoogle ScholarPubMed
Shi-Wen, X, Leask, A and Abraham, D 2008. Regulation and function of connective tissue growth factor/CCN2 in tissue repair, scarring and fibrosis. Cytokine & Growth Factor Reviews 19, 133144.CrossRefGoogle ScholarPubMed
Siegel, RC and Fu, JC 1976. Collagen cross-linking. Purification and substrate specificity of lysyl oxidase. Journal of Biological Chemistry 251, 57795785.Google ScholarPubMed
Siegel, RC, Fu, JC and Chang, Y 1976. Collagen cross-linking: the substrate specificity of lysyl oxidase. Advances in Experimental Medicine and Biology 74, 438446.CrossRefGoogle ScholarPubMed
Soderhall, C, Marenholz, I, Kerscher, T, Ruschendorf, F, Esparza-Gordillo, J, Worm, M, Gruber, C, Mayr, G, Albrecht, M, Rohde, K, Schulz, H, Wahn, U, Hubner, N and Lee, YA 2007. Variants in a novel epidermal collagen gene (COL29A1) are associated with atopic dermatitis. PLoS Biology 5, e242.CrossRefGoogle Scholar
Suwanabol, PA, Kent, KC and Liu, B 2011. TGF-beta and restenosis revisited: a Smad link. Journal of Surgical Research 167, 287297.CrossRefGoogle ScholarPubMed
Uezumi, A, Ikemoto-Uezumi, M and Tsuchida, K 2014. Roles of nonmyogenic mesenchymal progenitors in pathogenesis and regeneration of skeletal muscle. Frontiers in Physiology 5, 68.CrossRefGoogle ScholarPubMed
Uezumi, A, Fukada, S, Yamamoto, N, Takeda, S and Tsuchida, K 2010. Mesenchymal progenitors distinct from satellite cells contribute to ectopic fat cell formation in skeletal muscle. Nature Cell Biology 12, 143152.CrossRefGoogle ScholarPubMed
Uezumi, A, Ito, T, Morikawa, D, Shimizu, N, Yoneda, T, Segawa, M, Yamaguchi, M, Ogawa, R, Matev, MM, Miyagoe-Suzuki, Y, Takeda, S, Tsujikawa, K, Tsuchida, K, Yamamoto, H and Fukada, S 2011. Fibrosis and adipogenesis originate from a common mesenchymal progenitor in skeletal muscle. Journal of Cell Science 124, 36543664.CrossRefGoogle ScholarPubMed
Urciuolo, A, Quarta, M, Morbidoni, V, Gattazzo, F, Molon, S, Grumati, P, Montemurro, F, Tedesco, FS, Blaauw, B, Cossu, G, Vozzi, G, Rando, TA and Bonaldo, P 2013. Collagen VI regulates satellite cell self-renewal and muscle regeneration. Nature Communications 4, 1964.CrossRefGoogle ScholarPubMed
Veit, G, Kobbe, B, Keene, DR, Paulsson, M, Koch, M and Wagener, R 2006. Collagen XXVIII, a novel von Willebrand factor A domain-containing protein with many imperfections in the collagenous domain. Journal of Biological Chemistry 281, 34943504.CrossRefGoogle ScholarPubMed
Velleman, SG 1999. The role of the extracellular matrix in skeletal muscle development. Poultry Science 78, 778784.CrossRefGoogle ScholarPubMed
Velleman, SG 2007. Muscle development in the embryo and hatchling. Poultry Science 86, 10501054.CrossRefGoogle ScholarPubMed
Virag, JA, Rolle, ML, Reece, J, Hardouin, S, Feigl, EO and Murry, CE 2007. Fibroblast growth factor-2 regulates myocardial infarct repair: effects on cell proliferation, scar contraction, and ventricular function. American Journal of Pathology 171, 14311440.CrossRefGoogle ScholarPubMed
Voloshenyuk, TG, Hart, AD, Khoutorova, E and Gardner, JD 2011. TNF-alpha increases cardiac fibroblast lysyl oxidase expression through TGF-beta and PI3Kinase signaling pathways. Biochemical and Biophysical Research Communications 413, 370375.CrossRefGoogle ScholarPubMed
Wang, YX, Dumont, NA and Rudnicki, MA 2014. Muscle stem cells at a glance. Journal of Cell Science 127, 45434548.CrossRefGoogle ScholarPubMed
Wang, HT, Liu, CF, Tsai, TH, Chen, YL, Chang, HW, Tsai, CY, Leu, S, Zhen, YY, Chai, HT, Chung, SY, Chua, S, Yen, CH and Yip, HK 2012. Effect of obesity reduction on preservation of heart function and attenuation of left ventricular remodeling, oxidative stress and inflammation in obese mice. Journal of Translational Medicine 10, 145.CrossRefGoogle ScholarPubMed
Woessner, JF 1991. Matrix metalloproteinases and their inhibitors in connective tissue remodeling. The FASEB Journal 5, 21452154.Google ScholarPubMed
Wosczyna, MN, Biswas, AA, Cogswell, CA and Goldhamer, DJ 2012. Multipotent progenitors resident in the skeletal muscle interstitium exhibit robust BMP-dependent osteogenic activity and mediate heterotopic ossification. Journal of Bone and Mineral Research 27, 10041017.CrossRefGoogle ScholarPubMed
Yamaguchi, Y, Mann, DM and Ruoslahti, E 1990. Negative regulation of transforming growth factor-β by the proteoglycan decorin. Nature 346, 281284.CrossRefGoogle ScholarPubMed
Yang, QY, Liang, JF, Rogers, CJ, Zhao, JX, Zhu, MJ and Du, M 2013. Maternal obesity induces epigenetic modifications to facilitate zfp423 expression and enhance adipogenic differentiation in fetal mice. Diabetes 62, 37273735.CrossRefGoogle ScholarPubMed
Zhang, X, Nestor, KE, McFarland, DC and Velleman, SG 2008. The role of syndecan-4 and attached glycosaminoglycan chains on myogenic satellite cell growth. Matrix Biology 27, 619630.CrossRefGoogle ScholarPubMed
Zhang, X, Liu, C, Nestor, K, McFarland, D and Velleman, S 2007. The effect of glypican-1 glycosaminoglycan chains on turkey myogenic satellite cell proliferation, differentiation, and fibroblast growth factor 2 responsiveness. Poultry Science 86, 20202028.CrossRefGoogle ScholarPubMed
Zhao, T, Zhao, W, Chen, Y, Li, VS, Meng, W and Sun, Y 2013. Platelet-derived growth factor-D promotes fibrogenesis of cardiac fibroblasts. American Journal of Physiology – Heart and Circulatory Physiology 304, H1719H1726.CrossRefGoogle ScholarPubMed
Zhao, X, Wang, K, Liao, Y, Zeng, Q, Li, Y, Hu, F, Liu, Y, Meng, K, Qian, C, Zhang, Q, Guan, H, Feng, K, Zhou, Y, Du, Y and Chen, Z 2015. MicroRNA-101a inhibits cardiac fibrosis induced by hypoxia via targeting TGFbetaRI on cardiac fibroblasts. Cellular Physiology and Biochemistry 35, 213226.CrossRefGoogle ScholarPubMed
Zhou, B, Liu, Y, Kahn, M, Ann, DK, Han, A, Wang, H, Nguyen, C, Flodby, P, Zhong, Q, Krishnaveni, MS, Liebler, JM, Minoo, P, Crandall, ED and Borok, Z 2012. Interactions between beta-catenin and transforming growth factor-beta signaling pathways mediate epithelial-mesenchymal transition and are dependent on the transcriptional co-activator cAMP-response element-binding protein (CREB)-binding protein (CBP). Journal of Biological Chemistry 287, 70267038.CrossRefGoogle Scholar
Figure 0

Table 1 Factors enhancing and decreasing intramuscular fibrogenesis

Figure 1

Table 2 Growth factors associated with extracellular matrix and associated cells to regulate activation of satellite cells

You have Access