Skip to main content
PMC home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2011 Aug 1.
Published in final edited form as: J Neuropathol Exp Neurol. 2010 Aug;69(8):771–776. doi: 10.1097/NEN.0b013e3181e9a34b

Targeting Fibrosis in Duchenne Muscular Dystrophy

Lan Zhou 1, Haiyan Lu 1
PMCID: PMC2916968  NIHMSID: NIHMS221589  PMID: 20613637

Abstract

Duchenne muscular dystrophy (DMD) is the most common genetic muscle disease affecting 1 in 3,500 live male births. It is an X-linked recessive disease caused by a defective dystrophin gene. The disease is characterized by progressive limb weakness, respiratory and cardiac failure and premature death. Fibrosis is a prominent pathological feature of muscle biopsies from patients with DMD. It directly causes muscle dysfunction and contributes to the lethal DMD phenotype. Although gene therapy and cell therapy may ultimately provide a cure for DMD, currently the disease is devastating, with no effective therapies. Recent studies have demonstrated that ameliorating muscle fibrosis may represent a viable therapeutic approach for DMD. By reducing scar formation, antifibrotic therapies may not only improve muscle function but also enhance muscle regeneration and promote gene and stem cell engraftment. Antifibrotic therapy may serve as a necessary addition to gene and cell therapies to treat DMD in the future. Therefore, understanding cellular and molecular mechanisms underlying muscle fibrogenesis associated with dystrophin deficiency is key to the development of effective antifibrotic therapies for DMD.

Keywords: Antifibrotic therapy, Duchenne muscular dystrophy, Muscle fibrosis

INTRODUCTION

Fibrosis is defined as hardening and scar formation of tissues that results from uncontrolled wound-healing processes in response to chronic tissue injury and inflammation. It is characterized by excessive deposition of extracellular matrix (ECM) proteins, including collagens and fibronectin that can impair tissue function. Fibrosis can affect all tissues and organs causing considerable morbidity and mortality. Common fibrotic disorders include pulmonary fibrosis, cirrhosis, renal sclerosis, and scleroderma (1, 2).

Fibrosis is a prominent pathological feature of skeletal muscle in patients with Duchenne muscular dystrophy (DMD) (Fig. 1). DMD is the most common genetic muscle disease (3) and is characterized by progressive skeletal and cardiac muscle weakness with premature death usually around age 20 years (4). DMD is caused by a defective dystrophin gene on the X chromosome. Dystrophin deficiency disrupts the dystrophin-glycoprotein complex that normally spans muscle membranes to enable muscle to sustain mechanical stretch and contraction. A defective dystrophin-glycoprotein complex leads to increase of sarcolemmar permeability, influx of calcium into the sarcoplasm and activation of proteases to cause myofiber necrosis and degeneration. This in turn triggers an inflammatory response that may promote repair; the expression of the genetic defect results in chronic inflammation with persistent production of profibrotic cytokines and excessive synthesis and deposition of ECM proteins. A longitudinal study of 25 patients with DMD with a mean follow up of over 10 years showed that among the pathological features, including myofiber atrophy, necrosis, and fatty degeneration, only endomysial fibrosis on the initial muscle biopsies correlated with poor motor outcome gauged by muscle strength and age at loss of ambulation (5). This finding supports the notion that endomysial fibrosis directly contributes to progressive muscle dysfunction and the lethal phenotype of DMD.

Figure 1.

Figure 1

Open in a new tab

Collagen IV immunostaining shows that a muscle biopsy from a patient with Duchenne muscular dystrophy displays markedly increased endomysial collagen deposition (red) as compared to a normal muscle biopsy.

Currently there is no effective therapy for DMD. Gene therapy and cell therapy to replace the missing dystrophin gene have potential but are not yet sufficiently developed for widespread clinical application. The only relatively effective pharmacotherapy for DMD is corticosteroids, which prolongs independent ambulation by 2 to 4 years, but carries troublesome side effects. Although prednisone was initially evaluated to suppress muscle inflammation, its therapeutic mechanisms of action in DMD are still not entirely clear. At this point, there is no effective pharmacotherapy to attenuate muscle fibrosis in DMD patients.

Recent studies using the mdx mouse model of DMD have explored cellular and molecular mechanisms underlying skeletal muscle fibrogenesis associated with dystrophin deficiency and have tested several pharmacological agents to target muscle fibrogenesis. These studies provide compelling evidence that targeting muscle fibrosis can improve muscle function and the muscular dystrophy phenotype; therefore, this may represent a useful therapeutic approach for DMD.

RECENT RESULTS

Anti-fibrotic research in DMD has been mainly conducted on mdx mice. In this DMD model there is a nonsense mutation in exon 23 of the dystrophin gene and the mice display progressive endomysial fibrosis in diaphragm and cardiac muscles. These studies have largely focused on 3 aspects: 1) targeting signaling pathways of fibrogenic cytokines to inhibit ECM gene expression and protein synthesis; 2) suppressing muscle inflammation to reduce fibrogenic cytokine production; and 3) enhancing muscle regeneration (Table).

Table. Antifibrotic Interventions in mdx Mice.

Targeting Mechanisms Interventions Effects
Blocking fibrotic cytokine
signaling 		Decorin (TGF-β) ↓diaphragm collagen I mRNA
TGF-β neutralizing antibody ↓diaphragm fibrosis, ↑CD4+ T cells
Losartan (TGF-β) ↓diaphragm fibrosis
Halofuginone (TGF-β) ↓cardiac fibrosis
Fibrinogen depletion (TGF-β) ↓diaphragm fibrosis
Imatinib (TGF-β & PDGF) ↓diaphragm fibrosis
Myostatin−/−/mdx (TGF-β) ↓diaphragm fibrosis
Suppressing inflammation scid/mdx (lymphocytes) ↓diaphragm fibrosis
nu/nu/mdx (T lymphocyte) ↓diaphragm fibrosis
MBP-1−/−/mdx (eosinophils) ↓diaphragm fibrosis
Osteopontin−/−/mdx (Tregs) ↓diaphragm and cardiac fibrosis
Enhancing regeneration Mlgf+/+/mdx ↓diaphragm fibrosis
Myostatin−/−/mdx ↓diaphragm fibrosis
Open in a new tab

TGB-β = transforming growth factor-β.

Targeting Signaling Pathways of Fibrogenic Cytokines

ECM proteins are mainly produced by activated tissue fibroblasts. Activation of fibroblasts and expression of ECM proteins are stimulated by fibrogenic cytokines, including transforming growth factor-β1 (TGF-β1) and platelet-derived growth factor (PDGF), among others (1, 2).

TGF-β is the most potent fibrogenic cytokine and it contributes to the pathogenesis of a variety of fibrotic disorders (6-9), including muscular dystrophy (10). Tissue fibrosis is mainly regulated by the TGF-β1 isoform. It is an autocrine and paracrine cytokine that regulates fibrosis by signaling through its transmembrane serine/threonine kinase receptors of which there are 3 types. Type I (TβRI) and type II (TβRII) receptors are signaling receptors that form heterodimers, whereas the type III receptor (TβRIII) is a proteoglycan that regulates access of TGF-β to the signaling receptors. TGF-β binds directly to the TβRII subunit of the heterodimer, allowing TβRII to activate TβRI by phosphorylation. TβRI subsequently phosphorylates downstream Smad proteins, leading to translocation of the Smad complexes into the nucleus. Within the nucleus, the Smad complex binds to DNA in a sequence-specific manner to regulate transcription of many target genes, including fibrotic genes (11). TGF-β increases ECM deposition by stimulating synthesis of matrix proteins (12), reducing production of matrix-degrading proteases, and modulating expression of ECM receptors on the cell surface (13).

The expression patterns of TGF-β and its receptors in skeletal muscle of DMD patients and mdx mice support a pathogenic role for this fibrogenic cytokine in muscle fibrosis associated with dystrophin deficiency. TGF-β1 expression levels correlated with muscle fibrosis in muscle biopsies from patients with DMD (14). Moreover, expression of TGF-β and its receptors was also upregulated in skeletal muscles of mdx mice and were mainly localized to inflammatory and fibrotic areas (14, 15). Several studies have demonstrated that TGF-β signaling can be blocked by pharmacological or immunological means in mdx mice.

Decorin is a small leucine-rich proteoglycan that can bind TGF-β and inhibit its activity (16). Injection of decorin i.p. reduced collagen I mRNA expression in mdx diaphragm (17). Decorin also prevents differentiation of myogenic cells into fibrotic cells induced by TGF-β in injured skeletal muscles (18).

Andreetta et al tested whether immunomodulation of TGF-β1 could ameliorate diaphragm fibrosis in mdx mice (19). They treated mdx mice with i.p. injection of TGF-β1 neutralizing antibody from 6 to 12 weeks of age. The treated mdx mice showed significantly reduced diaphragm fibrosis along with decreased TGF-β1 mRNA and protein expression with no obvious effect on muscle degeneration or regeneration. However, because TGF-β1 is also an immunosuppressive cytokine, the treatment increased CD4+ lymphocytes. The authors suggested that long-term treatment with a TGF-β inhibitor should be evaluated for its overall effect on fibrosis and inflammation.

Losartan is an angiotensin II type 1 antagonist that is widely used as an antihypertensive medication. Angiotensin II directly stimulates TGF-β production and enhances TGF-β signaling by increasing Smad2 levels and the nuclear translocation of phosphorylated Smad3 (20). Thus, losartan can inhibit TGF-β signaling. Cohn et al treated mdx mice with oral losartan from 7 weeks to 9 months and showed that the treatment inhibited TGF-β signaling and significantly reduced mdx diaphragm fibrosis with no significant side effects (21).

Halofuginone is a potent antifibrotic agent that suppresses collagen synthesis mediated by the TGF-β signaling (22, 23) and inhibits phosphorylation and activation of Smad3 (24). Treatment of mdx mice with i.p. injection of halofuginone reduced the level of phosphorylated Smad3 and collagen deposition in limb and cardiac muscles; the latter was accompanied by improved cardiac function (25, 26). However, long-term use of halofuginone can cause skin tearing due to reduced collagen synthesis (27).

Fibrinogen is a soluble acute phase protein that extravasates at sites of inflammation to be converted to fibrin. Vidal et al found that fibrinogen deposition was increased in the fibrotic areas in DMD muscle biopsies and mdx diaphragm thereby implicating a role for this protein in muscle fibrogenesis. Indeed, mdx mice with fibrinogen deficiency showed reduced diaphragm fibrosis and treatment of mdx mice with ancrod, a defibrinogenating agent, reduced diaphragm fibrosis and degeneration, which was accompanied by decreased expression of TGF-β, phosphorylated Smad2, and collagen I. They further showed that fibrinogen could bind to the Mac-1 receptor on mdx macrophages to induce interleukin (IL)-1β expression and subsequent TGF-β synthesis, which in turn stimulated collagen expression by mdx fibroblasts. Fibrinogen also could bind to its αvβ3 integrin receptor on mdx fibroblasts to stimulate collagen synthesis directly (28). These findings indicate that targeting fibrinogen may represent a useful anti-fibrotic approach, and that its therapeutic function is exerted, at least in part, by inhibiting macrophage TGF-β production.

Imatinib is an anti-neoplastic agent that selectively and competitively blocks the ATP binding sites of several tyrosine kinases, including c-abl, c-kit, and PDGF receptors (29) and has been approved by the US Food and Drug Administration for treating several malignancies. Imatinib has also been shown to reduce tissue fibrosis via blocking c-abl and PDGF receptor signaling pathways in many experimental mouse models of fibrotic disorders, including pulmonary fibrosis (30, 31), cirrhosis (32), renal sclerosis (33), and skin fibrosis (34). The c-abl signaling represents an alternative non-Smad pathway that mediates the fibrogenic effect of TGF-β and c-abl kinase activity can also be activated by PDGF (31, 33). Because the gene and protein expression of TGF-β and PDGF (as well as of their receptors) was upregulated in inflammatory cells and regenerating fibers of skeletal muscles of DMD patients and mdx mice (14, 35-37), we tested whether imatinib could reduce mdx diaphragm fibrosis. We treated mdx mice with daily i.p. injections of imatinib from 8 to 14 weeks (38). The treatment greatly attenuated skeletal muscle necrosis, inflammation and diaphragm fibrosis, and improved muscle function. Reduced clinical disease was accompanied by inhibition of c-abl and PDGF receptor phosphorylation and suppression of tumor necrosis factor (TNF) and IL-1β expression. In another study, Bizario et al also showed that oral imatinib ameliorated muscle dystrophy in exercised mdx mice. However, the drug caused significant weight loss in the mice (39), which is a common but manageable side effect in humans.

Taken together, the above studies support the notion that TGF-β plays a key role in muscle fibrogenesis associated with dystrophinopathy. Targeting TGF-β and other fibrogenic cytokines (e.g. PDGF) may represent a useful therapeutic approach to inhibit muscle fibrogenesis in DMD patients.

Suppressing Muscle Inflammation

Chronic inflammation in DMD is secondary to continuous muscle degeneration and necrosis due to the expression of the dystrophin gene defect. Because increased collagen deposition is prominent in inflammatory areas of skeletal muscle in DMD, the inflammation is believed to contribute to muscle fibrogenesis. Inflammatory cells are major cellular sources of fibrogenic growth factors (as evidenced by the expression of TGF-β and PDGF), and their receptors are also upregulated and localized with inflammatory cells in muscles of DMD patient and mdx mice (14, 35-37).

Inflammatory cells in mdx skeletal muscle consist of lymphocytes, macrophages, neutrophils, eosinophils and mast cells (40-44). In addition to growth factors they also produce lymphokines that play critical roles in establishing proinflammatory, anti-inflammatory, or pro-fibrotic tissue environments. Tissue fibrosis is tightly regulated by the phenotype of T helper (TH) cell response (45). Fibrogenesis is strongly linked to TH2 CD4+ T cell responses, which involve IL-4, IL-5, and IL-13. TH1 CD4+ cells produce interferon-γ (IFN-γ) and IL-12 to promote tissue inflammation but may attenuate fibrosis. Studies of inflammatory cells in mdx mice have revealed several important immune mechanisms underlying muscle fibrogenesis associated with dystrophin deficiency.

To determine the role of lymphocytes in muscle dystrophy in mdx mice, scid/mdx mice that are deficient in functional T and B lymphocytes were generated (46). The scid/mdx mice showed less diaphragm fibrosis at 12 months and decreased levels of activated TGF-β1 protein in the muscle compared to mdx mice. A lack of functional T cells alone in nu/nu/mdx mice also led to reduced diaphragm fibrosis at 24 weeks (47). These findings support a pathogenic role for T cells in mdx diaphragm fibrogenesis and indicate that lymphocytes are an important source of TGF-β1. On the other hand, near complete postnatal depletion of circulating T cells in mdx mice by thymectomy at age 4 weeks, followed by anti-CD4 and/or -CD8 antibody treatment failed to improve diaphragm fibrosis at 24 weeks. These results suggest that early activation of T cells or resident T cells may play a critical role in promoting mdx diaphragm fibrosis (48).

Macrophages are numerous in skeletal muscle of DMD patients and mdx mice and are major sources of TGF-β1 and PDGF. Depletion of circulating macrophages by intraperitoneal injection of F4/80 antibody in mdx mice from age 1 to 4 weeks significantly suppressed leg muscle necrosis and degeneration, suggesting that macrophages contributed to muscle damage at the early stage of the disease (41). Long-term effects of macrophage depletion on mdx skeletal muscle dystrophy have not been studied.

Macrophages can display different functional phenotypes depending on the tissue cytokine environment (49, 50). Classically activated macrophages (M1) are activated by the TH-1 cytokines, IFN-γ and IL-12, and they express high levels of inducible nitric oxide synthase (iNOS). They increase the expression of TNF, IL-1β, nitric oxide, and major histocompatibility complex II molecules to promote tissue inflammation. Alternatively activated macrophages (M2) are activated by the TH-2 cytokines, IL-4 and IL-13 and express high levels of arginase but a low level of iNOS. These macrophages suppress TH-1 response and inhibit expression of TNF and IL-1β. The mannose receptor CD206 appears to be a relatively specific marker of this cell population. Because arginase can catalyze arginine into L-proline, a precursor molecule for collagen synthesis, M2 are thought to promote tissue fibrosis. However, a recent study showed that arginase-1-expressing macrophages suppressed TH-2 cytokine-driven inflammation and fibrosis in liver induced by Schistosoma mansoni infection (51). In this disease model, arginase-1-expressing macrophages suppressed CD4+ T cell proliferation and inhibited both TH-1 and TH-2 responses in an arginine-dependent manner. These findings indicate that M2 can function as suppressors of tissue inflammation and fibrosis. L-arginine also suppressed muscle inflammation in mdx mice. Intraperitoneal injection of L-arginine in mdx mice from 5 to 7 weeks suppressed muscle inflammation and reduced expression of inflammatory mediators, including IL-1β, TNF, IL-6, and NF-κB (52). The effect of L-arginine on mdx muscle fibrogenesis has not been addressed. Villalta et al studied functional subsets of macrophages in mdx mice and found that both M1 and M2 were present at 4 weeks in mdx quadriceps muscles that displayed prominent necrosis and inflammation (53). In vitro, M1 lysed muscle cells by nitric oxide-mediated mechanisms, whereas M2 inhibited this effect through the competition of arginase with iNOS for the common substrate of arginine. At 12 weeks, whereas the expression of both iNOS and arginase was reduced, IL-4 and IL-10 expression was increased, deactivating the M1 phenotype. IL-10 also activated the M2c phenotype, promoting satellite cell proliferation for muscle regeneration. These results suggest that a shift in macrophage phenotypes might contribute to muscle regeneration and resolution of inflammation in mdx quadriceps; the macrophage subsets and their roles in mdx diaphragm fibrogenesis have not been studied.

Eosinophils are increased in DMD muscle biopsies and in mdx skeletal muscles (43). Eosinophils can promote TH-2 responses by producing IL-4 and IL-10 to influence tissue fibrogenesis. Eosinophils also produce major basic protein-1 (MBP-1), which can attenuate cellular immune responses. MBP-1−/−/mdx mice showed reduced diaphragm fibrosis at 18 months with no change of macrophages or mRNA expression of iNOS, TNF, or IFN-γ at 4 weeks (54). These findings indicate that eosinophil-derived MBP-1 promoted mdx diaphragm fibrogenesis. Thus, MBP-1 may serve as a target for treating muscle fibrosis in DMD.

Recently, Vetrone et al demonstrated that osteopontin promoted mdx muscle fibrosis by modulating inflammation (55). Osteopontin is secreted by many types of cells, including mononuclear inflammatory cells and myofibers in DMD. The expression of osteopontin was upregulated in mdx blood and skeletal muscle. Genetic ablation of osteopontin interfered with muscle infiltration of neutrophils and NKT cells, but promoted T regulatory cell migration. The altered inflammatory response led to reduced TGF-β1 expression and attenuated diaphragm and cardiac muscle fibrosis in mdx mice (55). These findings suggest that osteopontin may also represent a promising therapeutic target in DMD.

Enhancing Muscle Regeneration

Fibrosis is a late-stage pathological change seen in many chronic myopathies and increasing muscle regenerative capacity may be able to reduce muscle fibrosis. This concept is supported by the following studies.

Insulin-like growth factor 1 (IGF-1) is a trophic factor for skeletal muscle, which promotes muscle regeneration and protein synthesis pathways. Exogenous expression of IGF-1 greatly increased muscle mass and force generation and reduced diaphragm fibrosis at 14 months in mdx mice (56).

Myostatin, a member of TGF-β family, negatively regulates skeletal muscle growth. Myostatin deficiency enhanced increased skeletal muscle mass, myofiber diameter, and strength; improved muscle regeneration was accompanied by reduced diaphragm fibrosis in mdx mice (57). A subsequent study by the same group showed that myostatin deficiency reduced muscle fibrosis not only by enhancing muscle regeneration but also by inhibiting muscle fibroblast growth and function (58). Myostatin directly stimulated proliferation of muscle fibroblasts by binding to its receptor, activin receptor IIB, which was expressed on muscle fibroblasts. It stimulated ECM protein synthesis in muscle fibroblasts by activation of Smad, p38 MAPK, and Akt pathways.

UNANSWERED QUESTIONS AND FUTURE DIRECTIONS

Muscle fibrosis has been increasingly recognized as a cause of muscle dysfunction in DMD (5), and as a barrier for muscle gene and stem cell delivery and engraftment (59, 60). Research on cellular and molecular mechanisms underlying muscle fibrogenesis and testing potential antifibrotic therapies has been active in the past several years but there are still many unanswered questions important to the future development of targeted antifibrotic therapies for DMD.

A striking feature of mdx mouse skeletal muscle is that diaphragm and limb muscles have different fates. Limb muscles show a near complete spontaneous resolution of inflammation with no significant endomysial fibrosis, whereas diaphragm displays persistent inflammation with progressive fibrosis (15, 61, 62). This feature suggests that muscle fibrogenesis associated with dystrophin deficiency probably involves interplay of different cell types and is influenced by the specific cellular and molecular tissue environment. The mechanisms by which the limb muscles are protected and diaphragm muscles are susceptible to fibrosis are not understood. There may be multiple factors including differences in inflammatory cells, cytokine profiles, tissue effector fibroblast properties, and/or myogenic cells. Understanding the fundamental differences contributing to the different fates of these 2 types of skeletal muscles in mdx mice may lead to identification of new targets for treating muscle fibrosis.

Muscle inflammation and fibrosis are likely linked in DMD. Individual inflammatory cell components have been studied to limited extent in mdx mice, especially with respect to diaphragm fibrogenesis (43, 46-48, 55). Interactions among different types of inflammatory cells and between inflammatory cells and tissue effector fibroblasts in limb and diaphragm muscles need to be characterized further to uncover the complex immune mechanisms underlying diaphragm fibrogenesis. This line of mechanistic studies will also direct future development of immune therapies targeting muscle fibrosis.

Muscle fibrosis is caused by excessive endomysial deposition of ECM proteins and results from disturbed balance of ECM protein synthesis and degradation. The studies using mdx mice have mainly focused on how to inhibit ECM protein synthesis. It may also be important to explore how to promote degradation of excessive ECM proteins to reduce scars. Because ECM proteins provide important tissue structural support, it will be challenging to prevent local scar formation by inhibiting collagen synthesis or melt scars by promoting collagen degradation without inducing systemic side effects, such as skin tearing as seen with the use of halofuginone (27).

Most of the interventions to test antifibrotic therapies in mdx mice started before the morphological onset of diaphragm fibrosis (19, 21, 28, 38, 39). It is unclear whether these identified and available agents can reverse muscle fibrosis. Because boys with DMD often develop significant limb muscle fibrosis at an early age, it would be more clinically relevant to develop and test a therapy that can reverse or slow down the progression of existing muscle fibrosis.

Because mdx mice display a mild phenotype with a near normal lifespan, disease progression in mdx mice is apparently different from that in DMD patients. Thus, findings in mdx mice may not be translated directly to DMD patients. Although myostatin deficiency showed a great therapeutic effect on mdx mice, Becker muscular dystrophy patients treated with a neutralizing antibody against myostatin (MYO-029) showed no improvement of muscle strength or function (63). More potent myostatin inhibitors are needed for treating dystrophinopathy patients. It is conceivable that treating DMD patients would be much more difficult than treating mdx mice and a combination of antifibrotic therapies targeting different aspects of muscle fibrogenesis will likely be required.

CONCLUSIONS

DMD is currently lethal and untreatable. Comprehensive approaches combining gene therapy, cell therapy, and pharmacotherapy will likely be required for an ultimate cure for this devastating disease. Clinical trials testing currently identified and available antifibrotic agents in DMD are expected in the near future; in particular, pharmacological agents have been approved by the FDA to treat human diseases. More studies characterizing muscle fibrogenesis associated with dystrophin deficiency and exploring potential safe and effective antifibrotic therapies are needed. Muscle fibrosis not only causes muscle dysfunction, but also impairs muscle regeneration and reduces gene and stem cell delivery and engraftment efficiency (59, 60). Therefore, antifibrotic therapy will represent a useful and necessary addition to gene and cell therapies to optimally treat DMD. A pharmacotherapy “cocktail” targeting different aspects of muscle fibrogenesis is likely needed to achieve the adequate anti-fibrotic effect in DMD.

Acknowledgments

Work supported by NIH K08 NS049346 (LZ) and MDA#91682 (LZ).

Footnotes

This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

REFERENCES

  • 1.Wynn TA. Common and unique mechanisms regulate fibrosis in various fibroproliferative diseases. J Clin Invest. 2007;117:524–9. doi: 10.1172/JCI31487. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Wynn TA. Cellular and molecular mechanisms of fibrosis. J Pathol. 2008;214:199–210. doi: 10.1002/path.2277. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Emery AE. Population frequencies of inherited neuromuscular diseases--a world survey. Neuromuscul Disord. 1991;1:19–29. doi: 10.1016/0960-8966(91)90039-u. [DOI] [PubMed] [Google Scholar]
  • 4.Emery AEH. Duchenne Muscular Dystrophy. Oxford University Press; Oxford: 1993. [Google Scholar]
  • 5.Desguerre I, Mayer M, Leturcq F, et al. Endomysial fibrosis in Duchenne muscular dystrophy: a marker of poor outcome associated with macrophage alternative activation. J Neuropathol Exp Neurol. 2009;68:762–73. doi: 10.1097/NEN.0b013e3181aa31c2. [DOI] [PubMed] [Google Scholar]
  • 6.Border WA, Noble NA, Yamamoto T, et al. Natural inhibitor of transforming growth factor-beta protects against scarring in experimental kidney disease. Nature. 1992;360:361–4. doi: 10.1038/360361a0. [DOI] [PubMed] [Google Scholar]
  • 7.Clouthier DE, Comerford SA, Hammer RE. Hepatic fibrosis, glomerulosclerosis, and a lipodystrophy-like syndrome in PEPCK-TGF-beta1 transgenic mice. J Clin Invest. 1997;100:2697–2713. doi: 10.1172/JCI119815. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Sato M, Muragaki Y, Saika S, et al. Targeted disruption of TGF-beta1/Smad3 signaling protects against renal tubulointerstitial fibrosis induced by unilateral ureteral obstruction. J Clin Invest. 2003;112:1486–94. doi: 10.1172/JCI19270. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Sime PJ, Xing Z, Graham FL, et al. Adenovector-mediated gene transfer of active transforming growth factor-beta1 induces prolonged severe fibrosis in rat lung. J Clin Invest. 1997;100:768–76. doi: 10.1172/JCI119590. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10.Heydemann A, Ceco E, Lim JE, et al. Latent TGF-beta-binding protein 4 modifies muscular dystrophy in mice. J Clin Invest. 2009;119:3703–12. doi: 10.1172/JCI39845. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Shi Y, Massague J. Mechanisms of TGF-beta signaling from cell membrane to the nucleus. Cell. 2003;113:685–700. doi: 10.1016/s0092-8674(03)00432-x. [DOI] [PubMed] [Google Scholar]
  • 12.Runyan CE, Schnaper HW, Poncelet AC. Smad3 and PKCdelta mediate TGF-beta1-induced collagen I expression in human mesangial cells. Am J Physiol Renal Physiol. 2003;285:F413–22. doi: 10.1152/ajprenal.00082.2003. [DOI] [PubMed] [Google Scholar]
  • 13.Border WA, Noble NA. Transforming growth factor beta in tissue fibrosis. N Engl J Med. 1994;331:1286–92. doi: 10.1056/NEJM199411103311907. [DOI] [PubMed] [Google Scholar]
  • 14.Bernasconi P, Torchiana E, Confalonieri P, et al. Expression of transforming growth factor-beta 1 in dystrophic patient muscles correlates with fibrosis. Pathogenetic role of a fibrogenic cytokine. J Clin Invest. 1995;96:1137–44. doi: 10.1172/JCI118101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Zhou L, Porter JD, Cheng G, et al. Temporal and spatial mRNA expression patterns of TGF-beta1, 2, 3 and TbetaRI, II, III in skeletal muscles of mdx mice. Neuromuscul Disord. 2006;16:32–8. doi: 10.1016/j.nmd.2005.09.009. [DOI] [PubMed] [Google Scholar]
  • 16.Reed CC, Iozzo RV. The role of decorin in collagen fibrillogenesis and skin homeostasis. Glycoconj J. 2002;19:249–55. doi: 10.1023/A:1025383913444. [DOI] [PubMed] [Google Scholar]
  • 17.Gosselin LE, Williams JE, Deering M, et al. Localization and early time course of TGF-beta1 mRNA expression in dystrophic muscle. Muscle Nerve. 2004;30:645–53. doi: 10.1002/mus.20150. [DOI] [PubMed] [Google Scholar]
  • 18.Li Y, Foster W, Deasy BM, et al. Transforming growth factor-beta1 induces the differentiation of myogenic cells into fibrotic cells in injured skeletal muscle: A key event in muscle fibrogenesis. Am J Pathol. 2004;164:1007–19. doi: 10.1016/s0002-9440(10)63188-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Andreetta F, Bernasconi P, Baggi F, et al. Immunomodulation of TGF-beta 1 in mdx mouse inhibits connective tissue proliferation in diaphragm but increases inflammatory response: implications for antifibrotic therapy. J Neuroimmunol. 2006;175:77–86. doi: 10.1016/j.jneuroim.2006.03.005. [DOI] [PubMed] [Google Scholar]
  • 20.Rosenkranz S. TGF-beta1 and angiotensin networking in cardiac remodeling. Cardiovasc Res. 2004;63:423–32. doi: 10.1016/j.cardiores.2004.04.030. [DOI] [PubMed] [Google Scholar]
  • 21.Cohn RD, van Erp C, Habashi JP, et al. Angiotensin II type 1 receptor blockade attenuates TGF-beta-induced failure of muscle regeneration in multiple myopathic states. Nat Med. 2007;13:204–10. doi: 10.1038/nm1536. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Granot I, Halevy O, Hurwitz S, et al. Halofuginone: an inhibitor of collagen type I synthesis. Biochim Biophys Acta. 1993;1156:107–12. doi: 10.1016/0304-4165(93)90123-p. [DOI] [PubMed] [Google Scholar]
  • 23.Halevy O, Nagler A, Levi-Schaffer F, et al. Inhibition of collagen type I synthesis by skin fibroblasts of graft versus host disease and scleroderma patients: effect of halofuginone. Biochem Pharmacol. 1996;52:1057–63. doi: 10.1016/0006-2952(96)00427-3. [DOI] [PubMed] [Google Scholar]
  • 24.McGaha TL, Phelps RG, Spiera H, et al. Halofuginone, an inhibitor of type-I collagen synthesis and skin sclerosis, blocks transforming-growth-factor-beta-mediated Smad3 activation in fibroblasts. J Invest Dermatol. 2002;118:461–70. doi: 10.1046/j.0022-202x.2001.01690.x. [DOI] [PubMed] [Google Scholar]
  • 25.Turgeman T, Hagai Y, Huebner K, et al. Prevention of muscle fibrosis and improvement in muscle performance in the mdx mouse by halofuginone. Neuromuscul Disord. 2008;18:857–68. doi: 10.1016/j.nmd.2008.06.386. [DOI] [PubMed] [Google Scholar]
  • 26.Huebner KD, Jassal DS, Halevy O, et al. Functional resolution of fibrosis in mdx mouse dystrophic heart and skeletal muscle by halofuginone. Am J Physiol Heart Circ Physiol. 2008;294:H1550–61. doi: 10.1152/ajpheart.01253.2007. [DOI] [PubMed] [Google Scholar]
  • 27.Granot I, Bartov I, Plavnik I, et al. Increased skin tearing in broilers and reduced collagen synthesis in skin in vivo and in vitro in response to the coccidiostat halofuginone. Poult Sci. 1991;70:1559–63. doi: 10.3382/ps.0701559. [DOI] [PubMed] [Google Scholar]
  • 28.Vidal B, Serrano AL, Tjwa M, et al. Fibrinogen drives dystrophic muscle fibrosis via a TGFbeta/alternative macrophage activation pathway. Genes Dev. 2008;22:1747–52. doi: 10.1101/gad.465908. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Druker BJ. Imatinib as a paradigm of targeted therapies. Adv Cancer Res. 2004;91:1–30. doi: 10.1016/S0065-230X(04)91001-9. [DOI] [PubMed] [Google Scholar]
  • 30.Abdollahi A, Li M, Ping G, et al. Inhibition of platelet-derived growth factor signaling attenuates pulmonary fibrosis. J Exp Med. 2005;201:925–35. doi: 10.1084/jem.20041393. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Daniels CE, Wilkes MC, Edens M, et al. Imatinib mesylate inhibits the profibrogenic activity of TGF-beta and prevents bleomycin-mediated lung fibrosis. J Clin Invest. 2004;114:1308–16. doi: 10.1172/JCI19603. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Yoshiji H, Noguchi R, Kuriyama S, et al. Imatinib mesylate (STI-571) attenuates liver fibrosis development in rats. Am J Physiol Gastrointest Liver Physiol. 2005;288:G907–13. doi: 10.1152/ajpgi.00420.2004. [DOI] [PubMed] [Google Scholar]
  • 33.Wang S, Wilkes MC, Leof EB, et al. Imatinib mesylate blocks a non-Smad TGF-beta pathway and reduces renal fibrogenesis in vivo. Faseb J. 2005;19:1–11. doi: 10.1096/fj.04-2370com. [DOI] [PubMed] [Google Scholar]
  • 34.Distler JH, Jungel A, Huber LC, et al. Imatinib mesylate reduces production of extracellular matrix and prevents development of experimental dermal fibrosis. Arthritis Rheum. 2007;56:311–22. doi: 10.1002/art.22314. [DOI] [PubMed] [Google Scholar]
  • 35.Tidball JG, Spencer MJ, St Pierre BA. PDGF-receptor concentration is elevated in regenerative muscle fibers in dystrophin-deficient muscle. Exp Cell Res. 1992;203:141–9. doi: 10.1016/0014-4827(92)90049-e. [DOI] [PubMed] [Google Scholar]
  • 36.Zhao Y, Haginoya K, Sun G, et al. Platelet-derived growth factor and its receptors are related to the progression of human muscular dystrophy: an immunohistochemical study. J Pathol. 2003;201:149–59. doi: 10.1002/path.1414. [DOI] [PubMed] [Google Scholar]
  • 37.Zhou L, Rafael-Fortney JA, Huang P, et al. Haploinsufficiency of utrophin gene worsens skeletal muscle inflammation and fibrosis in mdx mice. J Neurol Sci. 2008;264:106–11. doi: 10.1016/j.jns.2007.08.029. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Huang P, Zhao XS, Fields M, et al. Imatinib attenuates skeletal muscle dystrophy in mdx mice. FASEB J. 2009;23:2539–48. doi: 10.1096/fj.09-129833. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Bizario JC, Cerri DG, Rodrigues LC, et al. Imatinib mesylate ameliorates the dystrophic phenotype in exercised mdx mice. J Neuroimmunol. 2009;212:93–101. doi: 10.1016/j.jneuroim.2009.05.006. [DOI] [PubMed] [Google Scholar]
  • 40.Spencer MJ, Montecino-Rodriguez E, Dorshkind K, et al. Helper (CD4(+)) and cytotoxic (CD8(+)) T cells promote the pathology of dystrophin-deficient muscle. Clin Immunol. 2001;98:235–43. doi: 10.1006/clim.2000.4966. [DOI] [PubMed] [Google Scholar]
  • 41.Wehling M, Spencer MJ, Tidball JG. A nitric oxide synthase transgene ameliorates muscular dystrophy in mdx mice. J Cell Biol. 2001;155:123–31. doi: 10.1083/jcb.200105110. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Hodgetts S, Radley H, Davies M, et al. Reduced necrosis of dystrophic muscle by depletion of host neutrophils, or blocking TNFalpha function with Etanercept in mdx mice. Neuromuscul Disord. 2006;16:591–602. doi: 10.1016/j.nmd.2006.06.011. [DOI] [PubMed] [Google Scholar]
  • 43.Cai B, Spencer MJ, Nakamura G, et al. Eosinophilia of dystrophin-deficient muscle is promoted by perforin-mediated cytotoxicity by T cell effectors. Am J Pathol. 2000;156:1789–96. doi: 10.1016/S0002-9440(10)65050-X. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Gorospe JR, Tharp M, Demitsu T, et al. Dystrophin-deficient myofibers are vulnerable to mast cell granule-induced necrosis. Neuromuscul Disord. 1994;4:325–33. doi: 10.1016/0960-8966(94)90068-x. [DOI] [PubMed] [Google Scholar]
  • 45.Wynn TA. Fibrotic disease and the T(H)1/T(H)2 paradigm. Nat Rev Immunol. 2004;4:583–94. doi: 10.1038/nri1412. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Farini A, Meregalli M, Belicchi M, et al. T and B lymphocyte depletion has a marked effect on the fibrosis of dystrophic skeletal muscles in the scid/mdx mouse. J Pathol. 2007;213:229–38. doi: 10.1002/path.2213. [DOI] [PubMed] [Google Scholar]
  • 47.Morrison J, Lu QL, Pastoret C, et al. T-cell-dependent fibrosis in the mdx dystrophic mouse. Lab Invest. 2000;80:881–91. doi: 10.1038/labinvest.3780092. [DOI] [PubMed] [Google Scholar]
  • 48.Morrison J, Palmer DB, Cobbold S, et al. Effects of T-lymphocyte depletion on muscle fibrosis in the mdx mouse. Am J Pathol. 2005;166:1701–10. doi: 10.1016/S0002-9440(10)62480-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Gordon S. Alternative activation of macrophages. Nat Rev Immunol. 2003;3:23–35. doi: 10.1038/nri978. [DOI] [PubMed] [Google Scholar]
  • 50.Martinez FO, Helming L, Gordon S. Alternative activation of macrophages: An immunologic functional perspective. Annu Rev Immunol. 2009;27:451–83. doi: 10.1146/annurev.immunol.021908.132532. [DOI] [PubMed] [Google Scholar]
  • 51.Pesce JT, Ramalingam TR, Mentink-Kane MM, et al. Arginase-1-expressing macrophages suppress Th2 cytokine-driven inflammation and fibrosis. PLoS Pathog. 2009;5:e1000371. doi: 10.1371/journal.ppat.1000371. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Hnia K, Gayraud J, Hugon G, et al. L-arginine decreases inflammation and modulates the nuclear factor-kappaB/matrix metalloproteinase cascade in mdx muscle fibers. Am J Pathol. 2008;172:1509–19. doi: 10.2353/ajpath.2008.071009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Villalta SA, Nguyen HX, Deng B, et al. Shifts in macrophage phenotypes and macrophage competition for arginine metabolism affect the severity of muscle pathology in muscular dystrophy. Hum Mol Genet. 2009;18:482–96. doi: 10.1093/hmg/ddn376. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Wehling-Henricks M, Sokolow S, Lee JJ, et al. Major basic protein-1 promotes fibrosis of dystrophic muscle and attenuates the cellular immune response in muscular dystrophy. Hum Mol Genet. 2008;17:2280–92. doi: 10.1093/hmg/ddn129. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Vetrone SA, Montecino-Rodriguez E, Kudryashova E, et al. Osteopontin promotes fibrosis in dystrophic mouse muscle by modulating immune cell subsets and intramuscular TGF-beta. J Clin Invest. 2009;119:1583–94. doi: 10.1172/JCI37662. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56.Barton ER, Morris L, Musaro A, et al. Muscle-specific expression of insulin-like growth factor I counters muscle decline in mdx mice. J Cell Biol. 2002;157:137–48. doi: 10.1083/jcb.200108071. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 57.Wagner KR, McPherron AC, Winik N, et al. Loss of myostatin attenuates severity of muscular dystrophy in mdx mice. Ann Neurol. 2002;52:832–6. doi: 10.1002/ana.10385. [DOI] [PubMed] [Google Scholar]
  • 58.Li ZB, Kollias HD, Wagner KR. Myostatin directly regulates skeletal muscle fibrosis. J Biol Chem. 2008;283:19371–8. doi: 10.1074/jbc.M802585200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 59.Cordier L, Hack AA, Scott MO, et al. Rescue of skeletal muscles of gamma-sarcoglycan-deficient mice with adeno-associated virus-mediated gene transfer. Mol Ther. 2000;1:119–29. doi: 10.1006/mthe.1999.0019. [DOI] [PubMed] [Google Scholar]
  • 60.Gargioli C, Coletta M, De Grandis F, et al. PlGF-MMP-9-expressing cells restore microcirculation and efficacy of cell therapy in aged dystrophic muscle. Nat Med. 2008;14:973–8. doi: 10.1038/nm.1852. [DOI] [PubMed] [Google Scholar]
  • 61.Dupont-Versteegden EE, McCarter RJ. Differential expression of muscular dystrophy in diaphragm versus hindlimb muscles of mdx mice. Muscle Nerve. 1992;15:1105–10. doi: 10.1002/mus.880151008. [DOI] [PubMed] [Google Scholar]
  • 62.Stedman HH, Sweeney HL, Shrager JB, et al. The mdx mouse diaphragm reproduces the degenerative changes of Duchenne muscular dystrophy. Nature. 1991;352:536–9. doi: 10.1038/352536a0. [DOI] [PubMed] [Google Scholar]
  • 63.Wagner KR, Fleckenstein JL, Amato AA, et al. A phase I/IItrial of MYO-029 in adult subjects with muscular dystrophy. Ann Neurol. 2008;63:561–71. doi: 10.1002/ana.21338. [DOI] [PubMed] [Google Scholar]

RESOURCES