Development of Novel Strategies to Improve Skeletal Muscle Repair and Adaptation Following Eccentric Exercise

DEVELOPMENT OF NOVEL STRATEGIES TO IMPROVE SKELETAL MUSCLE REPAIR AND ADAPTATION FOLLOWING ECCENTRIC EXERCISE

BY KAI ZOU

DISSERTATION Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Kinesiology in the Graduate College of the University of Illinois at Urbana-Champaign, 2014

Urbana, Illinois

Doctoral Committee: Assistant Professor Marni D. Boppart, Chair Associate Professor Kenneth R. Wilund Professor Jeffrey A. Woods Professor Jie Chen

ABSTRACT Eccentric contractions, often required for participation in resistance training, as well as daily activities, can injure skeletal muscle. The repair of muscle damage is essential for effective remodeling, tissue maintenance, and initiation of beneficial adaptations post-exercise. We have previously demonstrated that transgenic overexpression of the α7BX2 integrin in skeletal muscle

(MCK: α7BX2; α7Tg) enhances muscle repair and the adaptive response following eccentric exercise. Recent studies have provided evidence that mesenchymal stem cells residing in skeletal muscle (mMSCs) contribute to repair following injury by secreting a variety of factors that are important for progenitor cell (satellite cell) activation. Our lab has also previously established that mMSC proliferation and quantity is increased following eccentric exercise in a manner dependent on the presence of the α7BX2 integrin. Preliminary cell culture experiments conducted in our lab suggest that mMSC paracrine factor gene expression is enhanced in the presence of laminin, an important component of the basal lamina that provides the microenvironment for muscle stem cells. Thus, the purpose of this study was to determine the extent to which mMSC and laminin-111 (LM-111) supplementation can enhance muscle repair and/or the adaptive response associated with eccentric exercise.

Lipophilic dye-labeled mMSCs (Sca-1+CD45-) isolated from α7Tg muscle post-exercise via FACS were intramuscularly injected into 3 month old WT recipient mice. Controls were injected with equal volume saline. Mice either remained sedentary or were subjected to eccentric exercise training on a treadmill (3x/wk) for two or four weeks following mMSCs transplantation.

Gastrocnemius and soleus complexes were collected 24 hours after the last bout of exercise to analyze indices of muscle repair and growth. In a separate study, natural mouse LM-111 was injected intramuscularly into 3 month old WT recipient mice one week prior to exercise.

Controls were injected with equal volume saline. Mice either remained sedentary or were

ii subjected to a single bout of eccentric exercise on a treadmill for 30 minutes. Gastronemius and soleus complexes were collected 24 hours post-exercise to analyze indices of muscle repair. In this study, we demonstrate that mMSC supplementation increases satellite cell activation (2-fold compared to all other groups), myonuclear content, myofiber hypertrophy, and hindlimb strength post-exercise. In addition, the presence of supplemental LM-111 markedly increased satellite cell proliferation and content post-exercise. Additional experiments suggest that modification of endogenous mMSC function, including upregulation of hepatocyte growth factor (HGF) gene expression, may be responsible for satellite activation with LM-111.

Therefore, these data suggest that mMSC and LM-111 supplementation provide therapeutic strategies to enhance repair following eccentric exercise. Such strategies may not only benefit healthy individuals seeking to accelerate recovery and athletic performance, but also populations that have a limited capacity for skeletal muscle regeneration.

iii

ACKNOWLEDGEMENTS

I would like to express my deepest gratitude to my advisor, Dr. Marni Boppart, for her excellent guidance, caring and patience throughout this research and also with my professional development. I would also like to thank Michael De Lisio for his valuable revision input on my manuscripts and Carmen Valero for teaching me many lab techniques patiently. Additionally, a special thank you to Tor Jensen for helping me sort the stem cells whenever I needed. The projects would not be successful without his help. Finally, I would also like to thank my committee members Jeffrey Wood, Kenneth Wilund and Jie Chen for their time on my preliminary and final exams and their helpful comments and suggestions throughout the dissertation process.

I would also like to thank my lab mates Heather Huntsman, Yair Pincu, Ziad

Mahmassani, and Michael Munroe for their assistance in my dissertation and all the help to my work throughout the last 4.5 years. In addition, I would like to thank my undergraduate assistants who spent countless hours on microscope and image analysis, and without their help this work would not have been possible.

I would also like to thank the American College of Sports Medicine for their financial support of these projects.

I would also like to thank my family for their love, support, and encouragement during my graduate study.

Finally, I want to say thank you to my wife Huimin, without whom my contribution to this project would not be possible. I also want to thank my lovely son Grant for making my life so special.

TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION ...... 1 1.1 Study Specific Aims ...... 4 1.2 Summary ...... 6 1.3 References ...... 7

CHAPTER 2: LITERATURE REVIEW ...... 10 2.1 Skeletal Muscle Injury ...... 10 2.2 Skeletal Muscle Repair ...... 10 2.3 Mesenchymal Stem Cells ...... 18 2.4 Integrins and Skeletal Muscle ...... 27 2.5 Laminins and Skeletal Muscle ...... 30 2.6 Conclusion ...... 33 2.7 References ...... 34

CHAPTER 3: MESENCHYMAL STEM CELLS REGULATE MUSCLE REPAIR AND ADAPTIVE RESPONSE TO ECCENTRIC EXERCISE 45 3.1 Abstract ...... 45 3.2 Introduction ...... 47 3.3 Methods...... 51 3.4 Results ...... 57 3.5 Discussion ...... 61 3.6 Acknowledgements ...... 66 3.7 Figures, Table and Figure Legends ...... 67 3.8 References ...... 74

CHAPTER 4: LAMININ-111 ENHANCES THE MYOGENIC RESPONSE TO ECCENTRIC EXERCISE IN YOUNG AND AGED MICE ...... 78 4.1 Abstract ...... 78 4.2 Introduction ...... 80 4.3 Methods...... 83 4.4 Results ...... 90 4.5 Discussion ...... 94 4.6 Figures, Table and Figure Legends ...... 102 4.7 References ...... 112

CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS ...... 117

APPENDIX A ...... 119

APPENDIX B ...... 128

APPENDIX C ...... 155

CHAPTER 1 INTRODUCTION

Skeletal muscle is a highly structured and intricately organized organ that provides the basis for human movement. Physical interaction between the essential myofibrillar proteins, actin and myosin, within each myofibril of a skeletal muscle cell can allow for the development of tension and the transfer of force to tendon and bone. While concentric (shortening) contractions are most commonly utilized for movement of peripheral limbs during physical activity, lengthening of the muscle can also occur during actin-myosin cross-bridge formation

(eccentric contraction). The need for the muscle to contract during lengthening is most evident during activities that require transfer of a load away from the body (weightlifting) or during downward locomotion to prevent imbalance and falling. While repeatedly shortening the muscle confers predominantly metabolic adaptations that provide resistance to fatigue, a single event of lengthening can cause damage to the ultrastructure and initiate a regenerative response that can influence muscle growth and strength (14, 26). The extent to which fiber damage and repair is prerequisite for the strength gains associated with eccentric exercise is not known. Regardless, unaccustomed eccentric exercise can be used as a physiological model to study the process of adult skeletal muscle regeneration, providing information that can inform the development of strategies to reduce loss of muscle with age, immobilization, and disease.

1 and within the myotendinous and neuromuscular junctions (2). The presence of force at these specific sites in muscle suggests an important role for the α71 integrin in enhancement of tissue adhesion and resistance to mechanical damage. The rapid upregulation of several isoforms of the

7 integrin subunit RNA and/or protein in two mouse models of muscular dystrophy (mdx-/-, mdx/utr-/-), along with healthy wild type (WT) mice in response to eccentric exercise, further substantiates a role for the integrin in protection from damage (4, 5, 7, 17). In addition, 7 integrin knockout mice are highly susceptible to sarcolemmal and ultrastructural damage (24), while mice that overexpress the α7BX2 integrin in muscle are protected from damage and reductions in force that occurs following eccentric exercise (21). Thus, upregulation of α7 integrin protein expression in muscle appears to be an early event necessary for conferring resilience to mechanical strain and maintenance of tissue mass.

Our lab subsequently established that transgenic overexpression of the α71 integrin

(α7Tg) in mouse muscle enhances repair and the adaptive response to eccentric exercise (21).

Specifically, we have determined that enhanced expression of the integrin in muscle can increase satellite cell number, new fiber synthesis, hypertrophic signaling, myofibrillar content, individual fiber and whole muscle cross sectional area, and gains in maximal isometric force following single or multiple bouts of downhill running (21, 31). Thus, in addition to allowing protection from damage, the α7 integrin may play an important role in the process of healing, maintenance of tissue structure, and enhanced growth following exercise. Despite these intriguing observations, the precise mechanisms that underlie α7 integrin-mediated benefits are not clear.

Mesenchymal stem cells (MSCs), also known as mesenchymal stromal cells, reside in the vascular niche of a variety of tissues and enhance tissue repair following injury (1, 3, 9, 11, 13,

18, 22, 30). Although MSCs are multipotent and have the potential to differentiate into a wide

2 variety of mesodermal tissue types, MSCs primarily contribute to tissue healing through the release of paracrine factors that stimulate activation of tissue-resident progenitor stem cells. In skeletal muscle, MSCs have some myogenic potential and can directly form fibers de novo (3,

10), yet most studies suggest that MSCs predominantly activate the endogenous progenitors, satellite cells, to participate in regeneration (3, 6, 18, 22). We recently established that Sca-

1+CD45- MSCs are increased in muscle (mMSCs) in a manner dependent on the presence of the

α7 integrin following an acute bout of eccentric exercise (30). Extraction and transplantation of these cells into WT mice post-exercise demonstrate an indirect role in activating satellite cells and forming new fibers following eccentric exercise (30). These data suggest that mMSCs may provide the basis for enhanced healing previously observed in α7Tg mice post-exercise. The extent to which these cells can influence long-term healing, growth, and function of muscle following repeated bouts of eccentric exercise is not known.

Laminins are important and biologically active extracellular matrix proteins found in the basal lamina of skeletal muscle fibers that provide an important scaffold necessary for tissue organization, maintenance, and survival (19, 29). Several members of the integrin family, including α71 integrin, serve as receptors for laminin on a variety of cell types (8). Among the fourteen known isoforms, laminin-111(α1, β1, γ1) (LM-111) is the predominant isoform present during embryonic skeletal muscle development and is the preferred ligand for the α7 integrin subunit (12, 23). Although LM-111 does not exist in healthy adult muscle, LM-111 can be purified from mouse Engelbreth-Holm-Swarm (EHS) tumors. Interestingly, systemic or intramuscular injection of EHS LM-111 can effectively increase α7 integrin subunit expression while also stimulating myoblast proliferation and new fiber formation in different mouse models of neuromuscular disease (15, 16, 20, 27). Despite the promise for LM-111 to be used as a

3 therapeutic intervention, the precise mechanism by which LM-111 can influence muscle regeneration is not clear.

The long-term goal of this study is to evaluate the ability for mMSCs and/or LM-111 to facilitate muscle repair and the adaptive response to eccentric exercise. Our central hypothesis is that injection of mMSCs and/or LM-111 in young mice will effectively enhance muscle repair and promote beneficial adaptations following eccentric exercise. A brief rationale, methods, hypotheses, expected results, and alternative hypotheses are presented below. A full description of the study methodology, including experimental design and statistical analysis, can be found in each chapter.

1.1 Study Specific Aims

Specific Aim 1: We will determine the extent to which mMSC transplantation can facilitate muscle repair and growth following eccentric exercise training.

In this study, Sca-1+CD45- cells will be isolated using fluorescence-activated cell sorting

(FACS). Three month old female C57Bl/6 mice will be injected with either saline or mMSCs one hour following a single bout of downhill running. Recipient mice will be subjected to a single bout of eccentric exercise prior to injection to recapitulate the exercised microenvironment. Two days after injection, mice will either remain sedentary (SED) or will be subjected to repeated bouts of eccentric exercise for two or four weeks (TR). Thus, the four groups compared will be:

Saline/SED, Saline/TR, mMSCs/SED and mMSCs/TR (n = 6/group). Twenty-four hours following the final training session, all mice will be euthanized. Gastrocnemius-soleus muscles will be rapidly dissected and either frozen in liquid nitrogen prior to protein extraction or frozen in liquid nitrogen-cooled isopentane for immunohistochemistry studies.

We hypothesize that mMSC transplantation will increase satellite cell accumulation and facilitate muscle repair after two weeks of eccentric exercise training compared to saline treated controls. We also predict that enhancement of satellite cell number will contribute to an elevation in myonuclear content, ultimately allowing for increased myofiber growth and muscle strength following four weeks of eccentric exercise training. Based on previous results, mMSC transplantation alone will not be sufficient to observe a change in satellite cell number or myofiber growth; rather mMSC transplantation concurrent with exercise will allow for these beneficial changes. In vitro experiments will be completed to demonstrate that mMSC paracrine factor release is dependent on mechanical stain and that this event may provide the basis for in vivo observations.

The possibility exists that mMSC transplantation will not allow for enhancement of repair and growth post-exercise. This observation would suggest that mMSCs are not capable of properly functioning or surviving in the exercised microenvironment, and therefore, do not contribute to repair or adaptation post-eccentric exercise.

2 Specific Aim 2: We will determine the extent to which LM-111 supplementation can facilitate muscle repair following an acute bout of eccentric exercise.

In this study, 3-month-old C57Bl/6 mice will be used. Mice will receive a single intramuscular injection of EHS LM-111(100 µL, 1mg/ml) in sterile TBS (SA, n=12) or sterile

TBS (100 µL) (n=12) in both gastrocnemius muscles (25). At one week post-injection, mice will be subjected to a single bout of downhill running exercise (-20o, 17 m/min, 60 min) or remain sedentary. Thus, the four groups will be: Saline/SED, SA/EX, LM-111/SED and LM-111/EX

(n=6/group). Twenty-four hours following eccentric exercise, all mice will be euthanized via

5 carbon dioxide asphyxiation. The gastrocnemius-soleus complexes will be rapidly dissected and either frozen in liquid nitrogen for isolation of protein, pre-cooled isopentane for immunohistochemistry studies, or directly used for FACS to isolate Sca-1+CD45- cells.

Based on previous results, we hypothesize that LM-111 supplementation will upregulate

α7 integrin protein expression and allow for an increase in satellite cell number in skeletal muscle following an acute bout of eccentric exercise. Alternatively, LM-111 injection may enhance repair post-exercise without any change in α7 integrin expression. Preliminary data collected in our lab suggest that mMSC paracrine factor gene expression is sensitive to extracellular matrix proteins that comprise the muscle microenvironment. Specifically, mMSC paracrine factor gene expression is enhanced in the presence of laminin compared to collagen. In addition, other studies have demonstrated that LM-111 can directly improve myoblast proliferation, differentiation and migration in vitro (16). Therefore, LM-111 supplementation could be a feasible strategy to manipulate gene expression and function of endogenous stem cells.

1.2 Summary

The overall goal of this project is to evaluate the ability for mMSC transplantation and

LM-111 to enhance repair of skeletal muscle following injury associated with eccentric exercise.

The regenerative response to contraction-mediated damage is decreased with age and may largely account for the significant loss of muscle mass over the lifespan. Therefore, these experiments provide the first steps in determining the potential for mMSC and LM-111-based therapies to effectively prevent or recover loss of muscle mass associated with age.

1.3 References

1. Ambrosio F, Ferrari RJ, Distefano G, Plassmeyer JM, Carvell GE, Deasy BM, Boninger ML, Fitzgerald GK, and Huard J. The synergistic effect of treadmill running on stem-cell transplantation to heal injured skeletal muscle. Tissue Eng Part A 16: 839-849, 2010. 2. Bao ZZ, Lakonishok M, Kaufman S, and Horwitz AF. Alpha 7 beta 1 integrin is a component of the myotendinous junction on skeletal muscle. J Cell Sci 106 ( Pt 2): 579-589, 1993. 3. Birbrair A, Zhang T, Wang ZM, Messi ML, Enikolopov GN, Mintz A, and Delbono O. Role of Pericytes in Skeletal Muscle Regeneration and Fat Accumulation. Stem Cells Dev 22: 2298-2314, 2013. 4. Boppart MD, Burkin DJ, and Kaufman SJ. alpha(7)beta(1)-integrin regulates mechanotransduction and prevents skeletal muscle injury. Am J Physiol Cell Physiol 290: C1660-C1665, 2006. 5. Boppart MD, Volker SE, Alexander N, Burkin DJ, and Kaufman SJ. Exercise promotes alpha 7 integrin gene transcription and protection of skeletal muscle. Am J Physiol Regul Integr Comp Physiol 295: R1623-R1630, 2008. 6. Boyle M, Wong C, Rocha M, and Jones DL. Decline in self-renewal factors contributes to aging of the stem cell niche in the Drosophila testis. Cell Stem Cell 1: 470-478, 2007. 7. Burkin DJ, Wallace GQ, Nicol KJ, Kaufman DJ, and Kaufman SJ. Enhanced expression of the alpha 7 beta 1 integrin reduces muscular dystrophy and restores viability in dystrophic mice. J Cell Biol 152: 1207-1218, 2001. 8. Collo G, Starr L, and Quaranta V. A new isoform of the laminin receptor integrin alpha 7 beta 1 is developmentally regulated in skeletal muscle. J Biol Chem 268: 19019-19024, 1993. 9. Crisan M, Yap S, Casteilla L, Chen CW, Corselli M, Park TS, Andriolo G, Sun B, Zheng B, Zhang L, Norotte C, Teng PN, Traas J, Schugar R, Deasy BM, Badylak S, Buhring HJ, Giacobino JP, Lazzari L, Huard J, and Peault B. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell 3: 301-313, 2008. 10. Dellavalle A, Maroli G, Covarello D, Azzoni E, Innocenzi A, Perani L, Antonini S, Sambasivan R, Brunelli S, Tajbakhsh S, and Cossu G. Pericytes resident in postnatal skeletal muscle differentiate into muscle fibres and generate satellite cells. Nat Commun 2: 499, 2011. 11. Dellavalle A, Sampaolesi M, Tonlorenzi R, Tagliafico E, Sacchetti B, Perani L, Innocenzi A, Galvez BG, Messina G, Morosetti R, Li S, Belicchi M, Peretti G, Chamberlain JS, Wright WE, Torrente Y, Ferrari S, Bianco P, and Cossu G. Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat Cell Biol 9: 255-267, 2007.

12. Dev KK, Nishimune A, Henley JM, and Nakanishi S. The protein kinase C alpha binding protein PICK1 interacts with short but not long form alternative splice variants of AMPA receptor subunits. Neuropharmacology 38: 635-644, 1999. 13. Doyle MJ, Zhou S, Tanaka KK, Pisconti A, Farina NH, Sorrentino BP, and Olwin BB. Abcg2 labels multiple cell types in skeletal muscle and participates in muscle regeneration. J Cell Biol 195: 147-163, 2011. 14. Ebbeling CB, and Clarkson PM. Exercise-induced muscle damage and adaptation. Sports Med 7: 207-234, 1989. 15. Foster RF, Thompson JM, and Kaufman SJ. A laminin substrate promotes myogenesis in rat skeletal muscle cultures: analysis of replication and development using antidesmin and anti-BrdUrd monoclonal antibodies. Dev Biol 122: 11-20, 1987. 16. Goudenege S, Lamarre Y, Dumont N, Rousseau J, Frenette J, Skuk D, and Tremblay JP. Laminin-111: a potential therapeutic agent for Duchenne muscular dystrophy. Mol Ther 18: 2155-2163, 2010. 17. Hodges BL, Hayashi YK, Nonaka I, Wang W, Arahata K, and Kaufman SJ. Altered expression of the alpha7beta1 integrin in human and murine muscular dystrophies. J Cell Sci 110 ( Pt 22): 2873-2881, 1997. 18. Joe AW, Yi L, Natarajan A, Le Grand F, So L, Wang J, Rudnicki MA, and Rossi FM. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nat Cell Biol 12: 153-163, 2010. 19. Kleinman HK, and Weeks BS. Laminin: structure, functions and receptors. Curr Opin Cell Biol 1: 964-967, 1989. 20. Kroll TG, Peters BP, Hustad CM, Jones PA, Killen PD, and Ruddon RW. Expression of laminin chains during myogenic differentiation. J Biol Chem 269: 9270-9277, 1994. 21. Lueders TN, Zou K, Huntsman HD, Meador B, Mahmassani Z, Abel M, Valero MC, Huey KA, and Boppart MD. The alpha(7)beta(1)-integrin accelerates fiber hypertrophy and myogenesis following a single bout of eccentric exercise. Am J Physiol Cell Physiol 301: C938-C946, 2011. 22. Motohashi N, Uezumi A, Yada E, Fukada S, Fukushima K, Imaizumi K, Miyagoe- Suzuki Y, and Takeda S. Muscle CD31(-) CD45(-) side population cells promote muscle regeneration by stimulating proliferation and migration of myoblasts. Am J Pathol 173: 781-791, 2008. 23. Nishiuchi R, Takagi J, Hayashi M, Ido H, Yagi Y, Sanzen N, Tsuji T, Yamada M, and Sekiguchi K. Ligand-binding specificities of laminin-binding integrins: a comprehensive survey of laminin-integrin interactions using recombinant alpha3beta1, alpha6beta1, alpha7beta1 and alpha6beta4 integrins. Matrix Biol 25: 189-197, 2006.

24. Rooney JE, Gurpur PB, Yablonka-Reuveni Z, and Burkin DJ. Laminin-111 restores regenerative capacity in a mouse model for alpha7 integrin congenital myopathy. Am J Pathol 174: 256-264, 2009. 25. Rooney JE, Knapp JR, Hodges BL, Wuebbles RD, and Burkin DJ. Laminin-111 protein therapy reduces muscle pathology and improves viability of a mouse model of merosin- deficient congenital muscular dystrophy. Am J Pathol 180: 1593-1602, 2012. 26. Schoenfeld BJ. Does exercise-induced muscle damage play a role in skeletal muscle hypertrophy? J Strength Cond Res 26: 1441-1453, 2012. 27. Silva-Barbosa SD, Butler-Browne GS, de Mello W, Riederer I, Di Santo JP, Savino W, and Mouly V. Human myoblast engraftment is improved in laminin-enriched microenvironment. Transplantation 85: 566-575, 2008. 28. Song WK, Wang W, Sato H, Bielser DA, and Kaufman SJ. Expression of alpha 7 integrin cytoplasmic domains during skeletal muscle development: alternate forms, conformational change, and homologies with serine/threonine kinases and tyrosine phosphatases. J Cell Sci 106 ( Pt 4): 1139-1152, 1993. 29. Timpl R, Rohde H, Robey PG, Rennard SI, Foidart JM, and Martin GR. Laminin--a glycoprotein from basement membranes. J Biol Chem 254: 9933-9937, 1979. 30. Valero MC, Huntsman HD, Liu J, Zou K, and Boppart MD. Eccentric exercise facilitates mesenchymal stem cell appearance in skeletal muscle. PLoS One 7: e29760, 2012. 31. Zou K, Meador BM, Johnson B, Huntsman HD, Mahmassani Z, Valero MC, Huey KA, and Boppart MD. The alpha(7)beta(1)-integrin increases muscle hypertrophy following multiple bouts of eccentric exercise. J Appl Physiol (1985) 111: 1134-1141, 2011.

CHAPTER 2

LITERATURE REVIEW

2.1 Skeletal Muscle Injury

Unaccustomed exercise, particularly eccentric exercise, can cause ultrastructural damage to skeletal muscle (25, 34). Although eccentric contractions are often involved in planned resistance exercise; they also commonly occur during activities of routine daily life such as lowering heavy items or walking downstairs. Skeletal muscle injury is accompanied by pain, soreness and stiffness, followed by a period of reduced activity (55).

According to published studies in the US, muscle injuries, predominantly muscle contusion and strain injuries, account for approximately 90% injuries occurring in sports (114).

Furthermore, muscle repair from each session of exercise is critical for subsequent performance and continued improvements in strength with training. Therefore, any supplement or intervention that can accelerate muscle repair, growth, and tissue vascularization is important and necessary during periods of rest and/or rehabilitation. Strategies to improve muscle repair include the use of NSAIDs (94, 119), yet results are not consistent in the literature regarding acceleration of muscle repair (112) and the detrimental effects of long-term treatment may limit the application of these strategies (12).

Aging is associated with greater susceptibility to exercise-induced muscle injury and impaired regeneration capacity, both which likely contribute to the loss of muscle mass and force and the state of frailty in the elderly (23, 27, 68). Therefore, there is a critical need to develop novel and effective strategies to improve muscle repair.

2.2 Skeletal Muscle Repair

2.2.1 General Overview

Skeletal muscle is the largest organ of human body, comprising 40% of body mass, and is the source of power for locomotion. Skeletal muscle has a robust ability to regenerate in response to injury caused by direct trauma (e.g., unaccustomed and/or intense exercise, blunt trauma).

Skeletal muscle repair involves multiple cellular and molecular events which require a variety of mononuclear cell populations, including inflammatory cells, satellite cells and non-satellite stem cells (4). In general, the process consists of four interrelated phases: degeneration, inflammation, regeneration and remodeling (Figure 2.1) (101). Inflammatory cells and muscle stem cells routinely communicate to most effectively influence regeneration. Given the fact that subtle muscle injury occurs during activities of daily living, continuous repair is essential for tissue maintenance. Thus, understanding the mechanisms responsible for healing can be informative in maintaining health throughout the lifespan and in designing therapeutic interventions to improve the rate and efficiency of skeletal muscle healing following injury.

2.2.2 Degeneration and Inflammation

Muscle injury induces necrosis of damaged myofibers, which can be initiated by the development of a lesion within the myofiber sarcolemma (120). Thus, sarcolemmal damage allows for the detection of muscle damage via increased plasma levels of muscle protein, such as creatine kinase (102), or the uptake of certain dyes, such as Evans Blue Dye, by the damaged myofibers (49). In addition, sarcolemma breakdown is accompanied by calcium efflux by the sarcolemma reticulum, which can activate calcium-dependent proteolysis and exacerbate tissue degeneration (3).

The degeneration of the myofiber initiates an innate immune response, characterized by neutrophil and macrophage infiltration (120). Neutrophils are the first invaders of muscle after muscle injury (120). Generally, neutrophil infiltration occurs rapidly within one hour of muscle

11 injury (37), and reaches a peak 24 hours post injury. While neutrophils can secrete large amounts of cytolytic and cytotoxic molecules that further damage the injured muscle, they also can rapidly release high concentrations of free radicals that facilitate the removal of cellular debris

(120). Nguyen and colleagues have found that ablation of neutrophils prolongs the removal of cellular debris and delays muscle regeneration, suggesting neutrophils are necessary for skeletal muscle repair (79).

Macrophages are minimally present in the epimysium of healthy skeletal muscle tissue

(16). However, following the neutrophil invasion into injured muscle, macrophages are activated and elevated by 12 to 24 hours after injury (120). It has been well demonstrated that macrophages play an important role in muscle repair not only by assisting neutrophils to remove the debris, but also recruiting and activating satellite cells to be involved in muscle regeneration

(120, 121). This bi-functional characteristic of macrophages is attributed to the existence of different classes of macrophages in skeletal muscle, known as M1 (classically activated, pro- inflammatory and phagocytic) and M2 (alternatively activated, pro-regenerative).

Previous studies have described and characterized both classes, including different profiles of cytokine secretion (121). While the M1 subtype expresses CD68 and is typically IL-

12high and IL-10low, M2 macrophages express CD163, CD206 and are typically IL-10high and

IL-12low (121).

Figure 2.1. Phases of skeletal muscle regeneration. Adapted from Carosio, 2011.

2.2.3 Regeneration

Skeletal muscle regeneration requires the activation of tissue-resident progenitors, or satellite cells. Upon injury, these cells are activated, proliferate, differentiate, and fuse to form multinucleated myofibers. Non-satellite, multipotent stem cells resident to muscle interstitium may also contribute to muscle regeneration (2, 10, 33, 56, 77, 92, 128).

Satellite Cells

Satellite cells are quiescent muscle precursor cells in adult muscle and located outside the muscle fiber sarcolemma and beneath the basement lamina which are characterized by expression of the nuclear marker Pax7 (paired box protein7) (110). They are small mononuclear progenitor cells with virtually no cytoplasm found in mature muscle. In undamaged muscle, the majority of satellite cells are quiescent; they neither differentiate nor undergo cell division.

However, following the damage of the myofiber, quiescent satellite cells are activated with the upregulation the expression of Pax7 and myogenic markers (Myf-5 and MyoD) as early as 12

13 hours post injury and subsequently enter the cell cycle and initiate proliferation as skeletal myoblasts (Figure 2.2) (24). At this point, satellite cells are often referred to as myogenic precursor cells (MPC), or adult myoblasts (131). MyoD has been shown as a critical myogenic regulatory factor in satellite cell differentiation and muscle regeneration. In vitro cell culture experiments have demonstrated that MyoD-/- primary cells are not able to form myotubes in differentiation medium (100). In addition, an in vivo study supported this finding with the observation of an accumulation of MPCs and a concurrent decrease in newly formed myofibers in MyoD-/- mice (71).

A fraction of satellite cells may maintain Pax7 expression and downregulate MyoD, eventually withdrawing from the cell cycle and returning to quiescence. However, another fraction of satellite cells maintain MyoD expression, downregulate Pax7 and commit to differentiation via upregulation in the expression of myogenin following proliferation. (29). The transition from cell proliferation to terminal differentiation requires the inhibition of cell division and up-regulation of myogenin and MRF4 (19). The differentiation phase is followed by fusion with damaged myofibers or fusion with other myoblasts to form new myofibers, which can be identified by their small size, triangular shape, hyperchromatic and enlarged central nuclei, and expression of embryonic myosin heavy chain (eMHC) (76).

Figure 2.2. Model of satellite cells in muscle regeneration. Muscle regeneration is characterized by the activation of quiescent satellite cells that express Pax7. Satellite cells then undergo proliferation, differentiation and fusing to new/pre-existing myofibers. Adapted from Scime, 2009

Numerous studies have provided evidence that satellite cells are indispensable for skeletal muscle regeneration. Pax7, a marker specific to both activated and quiescent satellite cells (30) has enabled researchers to study the role of satellite cells during muscle regeneration.

Relaix et al. found that satellite cells from Pax7-/- mice exhibit impaired proliferation and differentiation capacity (95). Other investigators also observed reduced myofiber size and an increase in the number of small caliber myofibers in Pax7-/- mice. Finally, all these aforementioned characteristics in Pax7-/- mice are associated with unsuccessful muscle regeneration in response to injury (61). More recently, a model for the conditional and local depletion of Pax7-expressing muscle cells in mice was established (tamoxifen-induced expression of diphtheria toxin A; Pax7 iCE/+:R26R DTA/+ or Pax7-DTA). By using this model, the researchers provided unequivocal evidence for an essential role of Pax7+ satellite cells in adult regenerative myogenesis with the findings of dramatic loss of myofibers, inflammation, fat infiltration and loss of muscle mass in Pax7 after strenuous exercise (104). Consistently, three different genetic models based on the conditional ablation of satellite cells using the Pax7-DTA mouse model resulted in failure to regenerate skeletal muscle following injury (64, 70, 78). In

15 addition, indirect depletion of satellite cells by inhibiting Notch signaling through simultaneous constitutive inactivation of both Notch target genes can impair the regenerative response to injury (41).

Although a consensus exists that satellite cells are indispensable for muscle regeneration

(64, 70, 78, 104), there is still an unresolved question as to whether satellite cells are obligatory for skeletal muscle hypertrophy. The strongest evidence to date for the requirement of satellite cells for muscle hypertrophy is from the Pax7 knockout mouse. It was found that the postnatal muscle growth was severely blunted (82). However, these results were challenged since recent work has shown that satellite cells in adult muscle are different from satellite cells at earlier periods of development since they do not require Pax7 gene expression (63). Also Pax7 knockout mice have severe maturation problems and high mortality (<10% survive to adulthood) (82).

Blocking DNA synthesis through the use of γ-irradiation or chemical agents has been widely used to address the necessity of satellite cells in skeletal muscle hypertrophy process which found either blunted or completed loss of the hypertrophic response to mechanical overload after the destruction of satellite cell populations. (38, 74, 99). Unfortunately, when these studies were published, there was not yet a well-established method to identify the satellite cells, so the effectiveness of satellite cell ablation by blocking DNA synthesis was unknown. A further limitation of these studies was the inability of γ-irradiation to ablate satellite cells specifically

(69), thus it is possible that other important cell populations for muscle hypertrophy were also depleted.

Results from a recent study using Pax7-DTA mice demonstrate that the hypertrophic response to a mechanical load is not altered (70). However, follow-up studies suggested growth cannot be sustained with long-term loading, suggesting the need for satellite cells to supplement

16 the growth process. In addition, the fact that only 90% of satellite cells are removed provides the opportunity for remaining cells to respond to mechanical stimuli and participate in the hypertrophic response.

Non-satellite stem cells

In addition to satellite cells, a number of non-satellite stem cells have been identified that contribute to repair of muscle in response to injury. The precise origin of these cells and the predominant mechanisms by which they influence tissue healing remain speculative (10, 29, 33,

56, 71, 77, 100, 101). For a more extensive and detailed review regarding the role for the non- satellite stem cells in repair, refer to the section on “Mesenchymal Stem Cells” below.

2.2.4 Exercise-Induced Skeletal Muscle Repair

Unaccustomed, intense or long-duration exercise can induce damage to skeletal muscle, an event that initiates repair and remodeling of the tissue (25, 34). Exercise-induced muscle damage (EIMD) commonly occurs following engagement in activity that requires eccentric contractions compared to concentric and isometric contractions (26). Eccentric contractions are integral to resistance exercise, but also may be present during activities of daily living such as lowering heavy items (grocery bags) or walking downstairs. Ultrastructural damage to the individual myofibers can cause an immediate reduction in maximal isometric force, a deficit which may be sustained in the absence of repair (34). Alternatively, a robust regenerative response has the capacity to remodel muscle in such a way to allow for an enhancement in growth and strength (108). The precise mechanisms that underlie improvements in growth and strength following eccentric exercise are not yet clear. Therefore, delineating the early events in muscle repair process that occur with eccentric exercise can be informative in understanding the adaptive response to eccentric exercise.

Chemical injury, rather than contraction-mediated injury, has historically provided the basis for our currently understanding of muscle repair in mice. While the overall events important for repair, including degeneration of myofibers, the inflammatory response, and proliferation and activation of satellite cells, has been observed in response to eccentric contractions in mice and humans (86-88, 108, 117), the response is appropriately less robust.

In summary, studies from both animals and humans have demonstrated that exercise- induced muscle repair encompasses similar events such as inflammation and regeneration involving satellite cell activation.

2.3 Mesenchymal Stem Cells

2.3.1 General Overview

Mesenchymal stem cells (MSCs), also known as mesenchymal stromal cells, were first discovered in 1968 by Friedenstein as an adherent fibroblast-like cell population in bone barrow

(40). Other investigators then extended this observation and subsequently determined that MSCs can be isolated from various tissues throughout the body including, but not limited to, adipose tissue, peripheral blood, and tendon (103, 130, 134). Interest in MSC-based therapies has rapidly grown for many reasons; including the capacity for differentiation into multiple lineages, ease of expansion in vitro, and the lack of immunogenic response with allotransplantation (21, 58).

Whether MSCs should be categorized as stem cells as proposed by Caplan (20) or stromal cells as they are belong to stroma is still under debate. Therefore, it has been proposed that the term

“multipotent mesenchymal stromal cells” be adopted in place of “mesenchymal stem cells” (52).

The identification of MSCs is still elusive as no single universal marker can be used for a pure isolation. In general, a group of cell surface markers are used to identify MSCs. In 2006, the

International Society of Cellular Therapy defined MSCs by the following three criteria (32):

(1) MSCs must be adherent to plastic under standard tissue culture conditions;

(2) MSCs must express certain cell surface markers such as CD73, CD90, and CD105, and lack expression of other markers including CD45, CD34, CD14, or CD11b, CD79α or CD19 and HLA-DR surface molecules;

(3) MSCs must have the capacity to differentiate into osteoblasts, adipocytes, and chondroblasts under in vitro conditions. It should be noted that these criteria are not entirely valid for all the species. For example, mouse MSCs differ from the human MSCs. Also, strain difference exists in mouse MSCs in terms of marker expression and their behavior in culture (90).

MSC therapy has been effective in the treatment of a variety of clinical conditions. MSCs can enhance repair and regeneration of tissue and has been utilized to treat spinal cord injuries

(67) and myocardial infarction (1, 22), among others. The precise mechanisms by which MSCs contribute to repair are not yet clear. However, recent studies have suggested that the primary mechanism is via the release of paracrine factors (36, 44, 46, 59, 107). Indeed MSCs have been demonstrated to synthesize and release factors including, but not limited to, vascular endothelial growth factor (VEGF), fibroblast growth factor-2 (FGF-2), hepatocyte growth factor (HGF) and insulin-like growth factor-1 (IGF-1) (43).

2.3.2 MSCs in Regeneration

Multipotent MSCs reside within the perivascular niche of a variety of tissues, directly repairing injured tissue, or indirectly facilitating regeneration by secreting numerous cytokines and growth factors that can stimulate resident progenitor stem cells (72). Skeletal muscle repair and regeneration in response to injury has traditionally been attributed solely to the satellite cell population. However, a variety of non-satellite stem cells recently identified in muscle also contribute to repair in response to damage (Figure 2.3).

Figure 2.3. Mesenchymal-like stem cells in skeletal muscle. Pericytes and Mesenchymal progenitors are examples of Sca-1+CD45- MSCs found in muscle that help repair tissue in response to injury. Adapted from Pannérec 2012.

These mononuclear cells have been isolated and categorized as multipotent muscle- derived stem cells (MDSC) (92, 93), side population (SP) cells (48, 75, 77, 126), mesoangioblasts (105), mesenchymal progenitors (125), fibro/adipogenic progenitors (FAPs)

(56), PW1+ interstitial cells (PICs) (73) and pericytes (105) based on method of extraction, localization within muscle, cellular function, and heterogeneous cell surface markers (Figure 2.3).

Despite the inability to clearly and efficiently distinguish among these different non-satellite mMSCs, stem cell antigen-1 (Sca-1)/Ly-6A is a commonly expressed murine glycosyl phosphatidylinositol-anchored cell surface protein that is used in combination with other cell surface markers to identify MSCs in muscle. The majority of isolated Sca-1+ non-satellite stem cells do not express the hematopoietic stem cell marker CD45, or satellite cell marker Pax7, and

20 vary in their ability to spontaneously differentiate into skeletal muscle, yet can readily fuse with myoblasts in co-cultures and/or can secrete factors that potently activate satellite cells in response to injury or disease (5, 30, 77). Although these cell populations meet the criteria regarding to the panel of cell surface markers, researchers hesitate to identify them as MSCs due to the stringent criteria necessary for this classification. Regardless, one common characteristic non-satellite stem cells share is that they all contribute to muscle repair in response to injury. A variety of non-satellite stem cells will be listed and reviewed in the following section. Only studies demonstrating their contribution to skeletal muscle will be reviewed.

Muscle-Derived Stem Cells (MDSCs)

MDSCs, identified by Huard’s group by preplate technique were purified from postnatal mouse skeletal muscles, and have been shown that they can effectively contribute to skeletal muscle repair (2, 92). These cells have the capacity to differentiate into multiple cell types both in vivo and in vitro, suggesting they are MSC-like stem cells. Further characterization by immunostaining and FACS suggests that MDSCs are CD34+CD56+Flk+CD45-Pax7-Desmin+/- and vimentin+/-. Despite the fact that MDSCs are likely heterogeneous, preplating provides the opportunity to isolate non-satellite stem cells in large quantities necessary for therapeutic applications.

Qu-Petersen and colleagues have found that MDSCs can enhance repair of dystrophic muscle following transplantation compared to early preplate cells, which are enriched in myogenic precursor cells (92). In addition, MDSCs have high self-renewal ability, and unlike satellite cells, are not rejected by the host immune system following transplantation.

It has been proposed that direct differentiation of these cells to myofibers is not a major determinant for successful tissue repair. In fact, donor cells become fibroblastic and contribute to

21 scar formation rather than incorporating into regenerating fibers. Despite their limited ability to directly contribute to muscle repair after injury, MDSCs can indirectly enhance recovery from injury by increasing angiogenesis and reducing scar formation (81). Interestingly, the level of

VEGF expression is highly correlated with increased vascularity, improved muscle regeneration, and strength at four weeks post transplantation (81). Thus, this research group concluded that

MDSCs largely facilitate repair via paracrine factor release.

Side Population (SP) Cells

SP cells, first identified in the bone marrow based on Hoechst 33342 dye exclusion, were reported to be present in muscle and contribute to both muscle and vascular regeneration following injury (5, 48). While the majority of muscle SP cells are CD31+ endothelial cells, a fraction of muscle-derived SP cells negative for CD31 (CD31-CD45-) were found to proliferate and contribute to new fiber formation in response to CTX injection (126). CD31-CD45- SP cells extracted from regenerating muscle not only expressed several MSC cell surface markers such as platelet-derived growth factor receptor α (PDGFRα), but also demonstrated the unique capacity to spontaneously differentiate into adipocytes or form osteogenic cells in the presence of osteogenic media in vitro. In addition, CD31-CD45- SP cells retrieved from injured muscle expressed angiogenic factors (e.g. angiopoietin-1 (ang-1) and VEGF) and tumor growth factor beta (TGF-β) antagonists (e.g. follistatin). Therefore, the importance of this study was the identification of a sub-fraction of SP cells that could act as tissue-resident MSCs, and contribute to muscle repair post-injury.

Despite the suggestion that CD31-CD45- SP cells could directly contribute to new fiber synthesis post-injury, such potential was subsequently found to be limited. Motohashi and colleagues determined that CD31-CD45- SP cells do not readily become muscle, but rather

22 enhance transplantation and proliferation of exogenously injected myoblasts, and increase growth of myoblast-engrafted fibers following CTX injection (77). Further gene expression profiling suggested that SP cells synthesize a wide variety of paracrine factors, including numerous factors known to promote muscle repair. Specifically, metalloproteinase-2 (MMP-2) was highlighted as one factor that could be released and promote myoblast migration following injury.

Doyle and colleagues recently evaluated SP cell fate using an inducible reporter for abcg2 ( Abcg2CreERT2 x Rosa26-LacZ mice) (33). LacZ+ cells accumulated in the interstitium of muscle, minimally fused with pre-existing fibers, and gave rise to a variety of cell types, including cells expressing stem cell antigen-1 (Sca-1) and the pericyte marker, PDGFRβ. Mice deficient in the expression of abcg2, thus lacking SP cells, displayed impaired regeneration following CTX injection.

Altogether, these studies suggest that SP cells, predominantly those that express PDGFRα and β and are negative for CD31 or CD45, are mesenchymal-like stem cells and/or pericytes which indirectly contribute to repair post-injury.

Mesenchymal Progenitors/FAPs

CD31-CD45- SP cells, as described above, were identified as mesenchymal progenitor cells based on the expression of cell surface markers and given their potential to differentiate into adipocytes and osteogenic cells (125). Despite some suggestion that CD31-CD45- SP cells could become myogenic, investigators were not able to demonstrate substantial contribution to new fiber formation in vitro or in vivo. Consistent with these results, studies using similar cells classified as fibro/adipogenic progenitors (FAPs) demonstrated little capacity for myogenesis and significant potential for adipogenesis (56). Thus, MSC populations are all proposed to be the

23 source of ectopic fat deposition, yet the ultimate fate of these cells appears to be highly dependent on the microenvironmental cues provided by the muscle, cues largely determined by presence or absence of disease. Studies support an indirect role for FAPs in the regeneration of muscle in response to glycerol-induced injury (56). Therefore, maintenance of multipotency and inhibition of differentiation may be important factors contributing to whole muscle health.

PW1+ Interstitial Cells (PICs)

PICs were first identified in the interstitium of early postnatal skeletal muscle and are characterized by expression of PW1, a gene strongly expressed in myogenic cells (73). Although both satellite cells and PICs express PW1 as well as Pax7, they are two distinct myogenic progenitor cells that reside in skeletal muscle (73, 84). While the majority of PICs are PDGFRα+ and similar to FAPs mentioned above, a small portion of PICs have myogenic potential (84).

Recent studies suggest that PICs can be divided into two sub-populations based on PDGFRα expression. While PDGFRα- PICs display myogenic potential in vitro, PDGFRα+ PICs express both brown and white fat markers and can form adipocytes in culture, suggesting this sub- population may represent mesenchymal progenitors/FAPs. The extent to which sub-populations of PICSs directly or indirectly contribute to repair of muscle in response to injury is not known.

Pericytes

Pericytes are characterized by their distinct morphology, localization within the basement membrane of vessels, and expression of a unique panel of cell surface markers (NG2, CD146,

PDGFRβ) (28). Investigators have begun to delineate two fractions of NG2+ pericytes in muscle: type 1 characterized by negative expression for nestin, positive expression for PDGFRα

(PDGFRβ+CD146+Sca-1+CD34+Pax7-), and type 2 characterized by positive expression for nestin and negative expression for PDGFRα (PDGFRβ+CD146+Sca-1-CD34-Pax7-) (10). While

24 both types are able to proliferate in response to glycerol or BaCl2-induced injury, type 1 pericytes give rise to adipogenic cells and type 2 become myogenic in culture. The extent to which type 1 pericytes represent the mesenchymal progenitor or FAPs described above is not known (56,

125). Regardless, both type-1 pericytes and FAPs have the potential to indirectly enhance myogenic progenitor differentiation (10, 56).

2.3.3 MSCs in Exercise-Induced Regeneration

The α7β1 integrin heterodimer is a transmembrane linkage protein that is highly expressed in skeletal muscle (116). Data from our laboratory suggest that a single bout of eccentric exercise can increase α7 integrin RNA and protein expression (14, 15). We subsequently demonstrated that a single bout of eccentric exercise can significantly enhance new fiber synthesis and satellite cell expansion in out α7Tg mice, which have transgenic overexpression of α7 integrin protein (8-fold high compared to WT counterparts) (66).

Unfortunately, the mechanism by which these beneficial events occur in α7Tg mice is still unknown.

Meanwhile, it is well known that MSCs are increased in response to exercise (62, 83).

Thus, it prompted us to assess MSCs in our WT and α7Tg mice in response to exercise.

Recently, a highly regenerative Sca-1+CD45-Pax7- non-satellite stem cell fraction has been identified in muscle. Therefore, we chose to assess the same population in our studies. First, we found that Sca-1+CD45-Pax7- cells were significantly increased in WT muscle 24 hours post- exercise compared to non-exercise controls (4.3% at rest to 9.4% post-exercise), and the total percentage was further enhanced with overexpression of the α7 integrin (8.7% at rest to 16.2%)

(128). Second, it was further characterized and identified that the majority of our Sca-1+CD45- cells isolated from α7Tg muscle post-exercise were pericytes with high expression of some

25 pericytes markers such as NG2, CD146, PDGFRβ. In addition, multi-lineage potential was established. Since these cells met the criteria for MSCs, including morphology, expression of

MSC markers, and multi-lineage potential, we refer to our cells as muscle-derived MSCs, or mMSCs to distinguish them from MSCs derived from other tissue types such as bone marrow and adipose.

The concurrent elevation of mMSCs content with the facilitation of muscle repair in our

α7Tg mice post a single bout of eccentric exercise indicates that these mMSCs may be actively involved in muscle repair following eccentric exercise. To prove this hypothesis, transplantation experiments have been conducted by injecting mMSCs isolated from pre-exercised α7Tg mice to

WT recipient mice which received an acute bout of eccentric exercise one hour prior to injection.

While mMSCs did not directly give rise to newly established myofibers or vessels, they indirectly contributed to satellite cell expansion, new fiber synthesis, and vascular growth one week post transplantation (53, 128). Thus, we speculate that our mMSCs represent mesenchymal progenitors, FAPs and/or type-1 pericytes previously described and provide a stromal role in tissue repair post-exercise (11, 56, 125). In order to further confirm whether or not they are the same type of cell population, future experiments should focus on testing their adipogenic potential both in vitro and in vivo by looking at their responses to pre-adipogenic environment induced by injury.

Furthermore, we have assessed a variety of cytokines and growth factors that can potentially be secreted by mMSCs, including IGF-1, IL-6, VEGF, HGF, and EGF, immediately after isolation from pre-exercised muscles in vitro by applying mechanical load with Flexcell system. The data suggest that our mMSCs secrete a lot of these paracrine factors, and are sensitive to mechanical cues including exercise. The extent to which treatment of mMSCs with

26 mechanical cues can alter the secretome and improve engraftment upon injection into a fibrotic or less accommodating environment needs further analysis.

2.4 Integrins and Skeletal Muscle

Integrins are heterodimeric, transmembrane receptors consisting of an α and a β chain that are non-covalently associated. Integrins form clusters and allow cells to adhere to the extracellular matrix which provides the opportunity for cellular stabilization, and the ability to transmit mechanical and chemical information from the outside to the inside of the cell (116). At least 18 α subunits and 8 β subunits have been characterized, resulting in the identification of 24 unique possible integrin heterodimers (54). Each integrin heterodimer has its own binding specificity, even though the major adhesive proteins of the matrix, such as fibronectin and laminin are recognized by multiple integrins.

2.4.1 α7β1 Integrin and Skeletal Muscle

The α7β1 integrin is highly expressed in adult skeletal muscle and is enriched at the myotendenous junctions (MTJs) (57). It provides an important link between laminin in the basement membrane and the inner actin cytoskeleton (9). It has been reported that LM is the sole

ECM ligand for the α7β1 integrin (35, 60, 115). Although there is only one α7 gene, intricate splicing mechanisms result in several α7 isoforms. Splicing of α7 transcripts produce alternative cytoplasmic (A and B) and extracellular domain variants (X1 and X2) which contributes to the functional diversity of the integrin (116). Congenital myopathies caused by mutations in the α7 gene (ITGA7) in humans confirm a critical role for α7 integrin in skeletal muscle (50). In contrast, enhanced expression of the α7β1 integrin in skeletal muscle of mice with a severe form of muscular dystrophy (mdx/utr-/-) slowed development of muscle pathology and markedly extends longevity (18).

2.4.2 α7β1 integrin and Skeletal Muscle Regeneration and Growth

The role of α7β1 integrin in muscle regeneration has been assessed using a mouse model for muscular dystrophy (mdx/utr−/− mice). Overexpression of α7 integrin in skeletal muscle

(α7BX2-mdx/utr−/−) lead to a two-fold increase in satellite cell number (17). Consistent with satellite cell data, the number of central located nuclei in five week old α7BX2-mdx/utr−/− transgenic mice was 1.7-fold greater than in nontransgenic mdx/utr−/− mice, suggesting improvements in regenerative capacity. Similar to these results, total α7B integrin protein expression is upregulated and can reinforce sarcolemmal integrity following an acute bout of eccentric exercise in healthy mice (14, 15). Overexpression of α7BX2 integrin in skeletal muscle protects it from damage, accelerates satellite cell accumulation, and stimulates new fiber synthesis in response to acute bout of eccentric exercise (66). To further investigate the role of

α7 integrin in muscle repair, Rooney and colleagues injected a cardiotoxin to induce damage in

α7 knockout mice (α7-/-). They found α7 integrin deficient muscle exhibited defective regeneration with a delay in satellite cell activation when compared with wild type (97). The precise mechanisms that underlie the role for the α7 integrin in protection from damage and enhanced repair is still unclear. However, both in vivo and in vitro experiments have determined that the α7 integrin can stimulate myoblast proliferation and survival (13, 65), suggesting the α7 integrin is an important regulator of skeletal muscle regeneration.

There are very few studies that have examined a role for integrins in skeletal muscle hypertrophy. Early studies on mice models of muscular dystrophy have demonstrated the importance of α7 integrin in skeletal muscle development (18, 51, 116). p70S6K phosphorylation is preferentially increased, and fiber hypertrophy has been observed in mdx/utr-/- mice overexpressing the α7 integrin transgene (17). These data from dystrophic mice suggest that α7

28 integrin may confer an enhanced growth response in healthy skeletal muscle. During the last several years, our lab has investigated a role for 7 integrin in skeletal muscle exercise-induced muscle hypertrophy. Boppart et al. previously demonstrated that eccentric exercise can increase

α7RNA transcripts, including both extracellular (X1 and X2) and intracellular (A and B) isoforms in skeletal muscle three hours post-exercise (15). Concomitant elevation in α7B integrin protein is observed 24 hours post-exercise in whole muscle lysate, and integration of this molecule at the sarcolemma protects muscle from exercise-induced muscle damage (14, 15).

Recently, Lueders et al. found that MCK-driven overexpression of the α7 integrin increases individual myofiber hypertrophy one week following a single bout of eccentric exercise, with increased mTOR phosphorylation occurring two days following exercise (66). We also subjected α7 integrin transgenic mice (α7Tg) and WT controls to repeated bouts of eccentric exercise for four weeks. We found that mTOR and p70s6k phosphorylation, myofibrillar protein content, myofiber CSA, whole muscle area, and maximal isometric force were all significantly elevated in α7Tg muscle compared to WT controls (133). These studies suggest that the function of the α7 integrin is not only to provide protection from damage, but to ensure an appropriate cellular response necessary for growth. However, the extent to which satellite cell expansion or new fiber synthesis contribute to the hypertrophic response to eccentric exercise is not known.

Interestingly, we have demonstrated that Sca-1+CD45- MSCS, predominantly pericytes, are increased in skeletal muscle (mMSCs) following an acute bout of eccentric exercise (128).

Surprisingly, mMSCs are minimally present in α7 integrin knockout mice in the sedentary state and increased in α7Tg mice in the sedentary state and post-exercise. The extent to which mMSCs influence the regenerative response to eccentric exercise and account for the robust adaptive responses observed in α7Tg mice in response to eccentric exercise is not clear.

2.5 Laminins and Skeletal Muscle

Skeletal muscle fibers are surrounded by a layer of extracellular matrix material called the basal lamina. The basal lamina is a thin scaffold of specific extracellular protein, predominantly collagen IV and laminin, necessary for fiber attachment to the surrounding connective tissue (collagen I). This specialized framework of extracellular matrix (ECM) also provides important cues necessary for satellite cell migration and function (6).

Laminins are heterotrimers comprised of an α-chain, a β-chain, and a γ-chain that intersect and form a cross-like structure. Whereas the three short arms of the LM structure are important for LM cross-linking, the long arm provides the basis for adhesion with integrins on cellular membranes which helps anchor cells to the membrane. Since several genetic variants of

α-, β- and γ-chains have been identified (6), LM molecules are named according to their chain composition. For example, LM-111 consists of α1, β1, and γ1. To date, fourteen other chain combinations have been identified in vivo. LMs serve a critical role in regulating cell differentiation, migration, adhesion, and survival (7).

2.5.1 Maintenance of Skeletal Muscle Organization and Integrity

In the mature skeletal muscle, LM-211 and LM-221 are the major laminins surrounding each muscle fiber. In order to support muscle integrity, these two laminins bind to each other in order to form a strong crosslink in the basal lamina (106). In addition, laminins 211 and 221 also bind to other matrix proteins, such as nidogens, which connect to collagen IV (118). Thus, with the network assembled by laminins themselves and laminins with other matrix proteins, the basal lamina is well formed and can structurally support muscle while communicating to fibers via integrins. Defective laminins can cause muscles to improperly form, leading to muscle diseases.

For instance, deficiency of Laminin-211 can result in merosin-deficient congenital muscular

30 dystrophy type 1A (MDC1A) both in human and mice, the most common form of congenital muscular dystrophy.

2.5.2 Myogenesis

Myogenesis is a complex multi-step process that is strongly influenced by ECM cues (42).

Laminin 1 enhances proliferation of myoblasts, stimulates their myoblast motility, and leads them to form myofibers (39). Merosin (laminin-2 and -4; alpha 2-beta 1/beta 2-gamma 1) is the predominant laminin in the skeletal muscle basement membrane. Early studies showed that merosin can induce myoblast fusion and stability, ultimately preventing apoptosis (127). In addition, laminin provides binding sites for proteoglycans, such as perlecan (123). The glycosaminoglycan chains of the proteoglycans, in turn, provide binding sites for growth factors, such as fibroblast growth factors and transforming growth factors, which are critical for myogenesis (8, 91).

Developmental studies have elucidated a role for laminin-111 in fetal growth. Laminin-

111 is widely expressed in the membrane surrounding murine somites(122). After initial myogenic events, two laminin isoforms, laminin-211 and laminin 511, stimulate primary myoblast fusion and form the basement membrane (85). Laminin 411 expression becomes dominant during secondary myoblast fusion (85). Laminin α2 chain is also present in the human fetus from around the seventh week of gestation, reaching maximum expression levels at week

21 (89, 111), while the laminin α4 chain is strongly expressed at week 16 (109).

Taken together, laminins are important regulators of myogenesis and are present throughout muscle development.

Myogenesis is a complex multi-step process that is strongly influenced by ECM cues (42).

An early study has reported that myogenesis was enhanced on laminin coated plate compared to

31 collagen by increasing myoblast proliferation in vitro (39). Recently, Siegel and colleagues observed that muscle primary cells were 4-fold more motile on laminin than on other substrates

(113). Taken together, these studies and others suggest that myogenesis is dependent on the presence of laminin. Specifically, while LM-111 enhances proliferation of myoblasts, stimulates their motility, and facilitates myotube formation (39), merosin (laminin-2 and -4; alpha 2-beta

1/beta 2-gamma 1), on the other hand, promotes myotube stability by preventing apoptosis (127).

In addition, laminin provides binding sites for proteoglycans, including the predominant muscle proteoglycan perlecan (123). The glycosaminoglycan chains of the proteoglycans, in turn, provide binding sites for growth factors such as fibroblast growth factors and transforming growth factors, which are critical for myogenesis (8, 91).

2.5.3 LM-111 and α7β1 integrin

Among the fourteen known LM isoforms, laminin-111(α1, β1, γ1) (LM-111) is the most widely studied. LM-111, the first member of LM family, was discovered in 1979 via isolation from Engelbreth-Holm-Swarm (EHS) tumors (124).

The relation between LM and LM receptors is thought to play a pivotal role for skeletal muscle function and integrity. Only two LM receptors, dystroglycan and α71 integrin, are present in adult skeletal muscle (132). Studies indicate that LM-111 is a preferred ligand for

α7β1 integrin (31, 80). In addition, in vitro experiments have determined that LM-111 treated primary myoblasts have a 1.7-fold higher expression of α7B integrin compared to PBS-treated controls (96). Rooney and colleagues have also reported that a single systemic injection of LM-

111 can significantly increase α7 integrin protein in mdx mice. Specifically, α7A and α7B integrin isoforms increased 1.6-fold and 2.6-fold with LM-111 injection, respectively. (96).

Hence, LM-111 is a preferred ligand for the α7 integrin and binding regulates its expression in skeletal muscle.

2.5.4 LM-111 Supplementation in Skeletal Muscle

LM-111 is expressed during embryonic development but is absent in postnatal normal and dystrophic skeletal muscle (47). During embryonic myogenesis, immature murine somites are surrounded by a LM-111-rich basement membrane. The presence of LM-111 during development suggests it has the potential influence repair and regeneration in adult skeletal muscle following injury. During the past five years, several studies have examined the ability for

LM-111 to recover function in mouse models of neuromuscular disease (45, 96-98, 129). The results from these studies suggest that intramuscular injection of LM-111 can effectively restore muscle repair, improve isometric force and/or extend life in both mdx mice and the mouse model for LM-211 deficiency, MDC1A (45, 96-98, 129). Upregulation of α7 integrin protein expression and/or increased adhesion to the 7 integrin may provide a feasible explanation for these beneficial changes (17, 18), yet further studies are necessary verify these results.

2.6 Conclusion

In conclusion, skeletal muscle repair is necessary for the maintenance of skeletal muscle mass and function following unaccustomed eccentric exercise. In addition, a robust regenerative response may underlie the beneficial gains in mass and strength associated with repeated bouts of eccentric exercise. This study will examine the extent to which mMSCs and LM-111 can enhance repair and/or the adaptive response to eccentric exercise. This work is important for developing strategies to optimally preserve and restore muscle mass over the lifespan.

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CHAPTER 3

MESENCHYMAL STEM CELLS REGULATE MUSCLE REPAIR AND ADAPTIVE

RESPONSE TO ECCENTRIC EXERCISE

3.1 ABSTRACT

PURPOSE: Skeletal muscle repair is essential for effective remodeling, tissue maintenance, and initiation of beneficial adaptations post-eccentric exercise. Non-satellite stem cells, expressing mesenchymal stem cell (MSC) markers, accumulate in muscle following eccentric exercise and may contribute to healing and positive outcomes associated with training. The purpose of this study was to determine the extent to which MSCs isolated from skeletal muscle (mMSCs) facilitate long-term repair and growth following eccentric exercise training.

METHODS: Lipophilic dye-labeled mMSCs (Sca-1+CD45-) or saline were intramuscularly injected into 3 month old WT recipient mice. Mice either remained sedentary (SED) or were subjected to eccentric exercise training (TR) on a treadmill (3x/wk) for two or four weeks following transplantation. Gastronemius and soleus complexes were collected 24 hours after the last bout of exercise.

RESULTS: Dye-labeled mMSCs did not directly fuse with existing fibers. However, mMSC transplantation decreased time to recovery during eccentric exercise training. Specifically, macrophage content and number of large myofibers with centrally located nuclei were significantly lower in mMSC/TR compared to Saline/TR at two weeks. Satellite cell number was

2-fold higher in mMSC/TR compared to all groups at two weeks. The number of nuclei per fiber was elevated in mMSC/TR at two and four weeks. Mean CSA, percentage of larger caliber fibers

(>3000 µm2) and grip strength were significantly increased in mMSC/TR compared to

Saline/SED and mMSC/SED at four weeks.

CONCLUSION: The results from this study demonstrate that mMSC transplantation can indirectly enhance rate of healing, growth and strength following repeated bouts of eccentric exercise. Therefore, MSCs that naturally accumulate in skeletal muscle following eccentric contractions may provide the mechanistic basis for the adaptive response to exercise.

Keywords: α7 integrin, satellite cells, stem cell antigen-1 (Sca-1), downhill running, repair, growth

3.2 INTRODUCTION

Downhill running has been used as a model to investigate the natural processes of skeletal muscle injury and repair following eccentric exercise (2, 29, 30). Similar to injury created by other methods, such as cardiotoxin and crush injury, a single bout of downhill running can impair sarcolemmal integrity, evoke an immune response, and reduce maximal isometric force within 24 hours post-exercise. Within two to four days, satellite cell number is increased and newly formed fibers with centrally-located nuclei are apparent throughout the muscle.

Although previous studies suggest that repeated bouts of downhill running do not exacerbate muscle damage or lengthen the recovery process (36), few studies to our knowledge have evaluated the long-term adaptive response to downhill running.

The α71 integrin is a critical transmembrane protein that links laminin in the basement membrane with the actin cytoskeleton across the sarcolemma (3, 8). The increase in adhesion afforded by muscle-specific transgenic expression of α7BX2 integrin can enhance resistance to muscle fiber injury and subsequently ameliorate pathology in mice with a severe form of muscular dystrophy (mdx/utr-/-) (9). Similarly, our previous work has established that total 7B integrin protein expression is upregulated and can reinforce sarcolemmal integrity following an acute bout of eccentric exercise in healthy mice (5, 6). We subsequently demonstrated that transgenic expression of the α7BX2 integrin in skeletal muscle (MCK:α7BX2; α7Tg) accelerates satellite cell accumulation, new fiber synthesis, hypertrophic signaling, myofibrillar content, individual fiber and whole muscle cross sectional area, and maximal isometric force following either single or multiple bouts of downhill running (28, 45). The precise mechanisms by which this important adhesion molecule can positively regulate repair and growth of muscle fibers following exercise have not been fully established.

Skeletal muscle repair is a well-studied biological process requiring the coordinated recruitment of immune cells and activation of progenitor cells surrounding each myofiber.

Satellite cells, located in the niche between the sarcolemma and the basal lamina, can be activated (identified by myoD and myf5 expression), proliferate and upregulate genes necessary for myogenic differentiation (myogenin and mrf4 expression) in response to injury, including exercise-induced injury (26). Upon differentiation, satellite cells can fuse with damaged myofibers or fuse together to establish new fibers within muscle tissue (10). Studies that implement methods which allow for the temporal ablation of satellite cells in adult skeletal muscle have established an essential role for the satellite cell in repair following chemical injury

48 or mechanical loading (32, 37, 38). Thus, current research is focused on delineating the factors within the microenvironment that can regulate progenitor cell proliferation and function.

Non-satellite stem cells have been identified in skeletal muscle that can significantly enhance repair post-injury. Despite lack of consensus regarding nomenclature and classification, the majority of Pax7- mononuclear cells in murine skeletal muscle express stem cell antigen-1

(Sca-1) and a variety of MSC markers, including CD90 and platelet-derived growth factor receptor α (PDGFRα) (4, 12, 25, 34, 41). Non-satellite stem cells in muscle are uniquely isolated and categorized as side population (SP) cells, pericytes, mesenchymal progenitors, fibroadipogenic progenitors (FAPs), and PW1+ interstitial cells (PICs) (4, 14, 25, 34, 35, 41).

While some investigators have demonstrated that non-satellite cells have some potential for myogenesis (4, 13), the majority suggest an indirect and paracrine factor contribution to muscle healing (1, 25, 34, 35, 41). The extent to which non-satellite stem cells contribute to post- exercise repair has not been thoroughly evaluated.

Our laboratory recently established that Sca-1+CD45- MSCs increase in muscle (mMSCs) in an 7 integrin-dependent manner and contribute to satellite cell accumulation and new fiber synthesis one week following a single bout of eccentric exercise (42). Although some heterogeneity exists in the population of cells captured by positive selection for Sca-1 and negative selection for CD45 (hematopoietic stem cell marker), we were able to verify by FACS analysis following short-term maintenance and expansion in culture that these cells do not express satellite cell markers, yet highly express mesenchymal and pericyte markers and remain multipotent (42). Similar to the majority of studies examining a role for MSCs in the muscle repair process following cardiotoxin injection (15, 35, 41), mMSCs did not directly contribute to

49 new fiber formation or fuse with pre-existing fibers, but rather indirectly influenced satellite cell number and new fiber synthesis following an acute bout of exercise (42).

The extent to which mMSCs account for acceleration of muscle repair as well as beneficial adaptations in fiber size and strength previously observed in α7Tg muscle in response to acute eccentric exercise and exercise training (28, 45) has not been determined. Specific ablation of mMSCs, whereas a good idea, is not possible given the lack of consensus on whether distinct non-satellite stem cell fractions exist in muscle and/or whether these populations stably express one marker necessary for cell survival and/or function. In this study, we assess indices of post-exercise repair, growth, and function in WT mice supplemented with MSCs isolated from

α7Tg muscle. We hypothesized that mMSCs could facilitate long-term repair and beneficial adaptations following repeated bouts of eccentric exercise.

3.3 METHODS

Animals. Protocols for animal use were approved by the Institutional Animal Care and Use

Committee (IACUC) of the University of Illinois at Urbana-Champaign. Three month old female

C57/BL6 mice were purchased for injection of saline or mMSC transplantation (Charles River,

Wilmington, MA). Transgenic mice overexpressing the α7BX2 integrin in skeletal muscle

(SJ6/C57BL6; MCK: α7BX2; α7Tg) were used for extraction and isolation of mMSCs.

Transgenic mice were produced at the University of Illinois Transgenic Animal Facility as described (5, 9). All mice maintained in a temperature-controlled animal room maintained on a

12:12 light-dark cycle at the University of Illinois. Mice were fed standard laboratory chow and water ad libitum.

Isolation of Sca-1+CD45- Cells from Skeletal Muscle. Sca-1+CD45- cells were isolated via fluorescence activated cell sorting (FACS), as previously described (42). Briefly, gastrocnemius- soleus complexes were harvested from 5 week old 7Tg mice 24 hours following an acute bout of downhill running exercise as described below. After enzymatic digestion of the muscle tissue, filtered samples were incubated on ice with anti-mouse CD16/CD32 (1 μg/106 cells)

(eBioscience, San Diego, CA) for 10 min in order to block non-specific Fc-mediated interactions. Cells were stained in a cocktail of monoclonal anti-mouse antibodies Sca-1-PE

(600ng/106 cells) and CD45-APC (300 ng/106 cells) (eBioscience, San Diego, CA) diluted in 2%

FBS in PBS. FACS was performed using an iCyt Reflection System, located at Carle Hospital

(Urbana, IL). Negative and single-stained controls were used to establish gates. Sca-1+CD45- cells were collected in medium for culture (High glucose Dulbecco’s Modified Eagle’s Medium

(DMEM), 10% FBS, 5 μg/mL gentamycin), and seeded onto uncoated plastic tissue culture

4 2 dishes at 2.5 x 10 cells/cm . Cultures were incubated at 37°C and 5% CO2. Media was replaced

51 every 3 days until the cells reached 80-90% confluency. On average, cells were grown in culture for 7 days and were not passaged during this time. Cells were then labeled with lipophilic dye

(DiI) prior to injection.

Eccentric Exercise Training. All mice were subjected to a single bout of downhill running exercise (-20°, 17 m/min, 30 min) one hour prior to injection of either saline or mMSCs. Mice then remained sedentary or were subjected to multiple bouts of eccentric exercise. Exercise training consisted of downhill running 3 times/week (Monday, Wednesday and Friday) for two or four weeks (-20°, 17 m/min, 30 min) on a treadmill (Exer-6M, Columbus Instruments). For each exercise bout, the speed was gradually increased from 10 to 17 m/min over the first 2 minutes as a warm-up period. Due to the limited number of mMSCs that we can be isolate from muscle, we were unable to inject all animals to finish both exercise training studies at the same time. Therefore, two studies (two weeks and four weeks) were performed separately at different times in order to assess long-term muscle repair (two weeks) and adaptations (four weeks) respectively. mMSC Transplantation. One hour after downhill running, animals were anesthetized with 2% isofluorane, administered by inhalation. For the two week study, the right gastrocnemius muscle of each animal was transplanted with 4 x 104 cells suspended in 50 l HBSS (mMSC), whereas the left gastrocnemius muscle was transplanted with an equal volume of HBSS solution (Saline).

For mice exercising for four weeks, both legs were transplanted with either cells or saline in order to measure grip strength. For both the two and four week studies, animals were randomly assigned to either a sedentary (SED) or exercise training (TR) group. Thus, the four groups compared were: Saline/SED, Saline/TR, mMSC/SED and mMSC/TR (n = 5-6/group).

Tissue Collection and Preservation. Twenty-four hours following the final training session, all mice were euthanized via carbon dioxide asphyxiation. Gastrocnemius-soleus muscles were rapidly dissected and either frozen in liquid nitrogen prior to protein extraction or frozen in liquid nitrogen-cooled isopentane for immunohistochemistry studies.

Immunohistochemistry. Muscle complexes were divided at the midline along the axial plane. The distal end was embedded in OCT (Tissue-Tek; Fischer Scientific). Three transverse cryosections per sample (8μm non-serial sections, each separated by a minimum of 40 μm) were cut for each histological assessment using a CM3050S cryostat (Lecia, Wezlar, Germany). Sections were placed on microscope slides (Superfrost; Fischer Scientific, Hanover Park, IL) and stored at -

80°C before staining. Sections were stained with antibodies against dystrophin (1:100, Abcam,

Cambridge, MA); Pax7 (1:10, DSHB, Iowa City, Iowa) and eMHC (1:8, BF-45, DSHB, Iowa

City, Iowa). All species-appropriate secondary antibodies used for immunofluorescence studies were applied at 1:200 (Pax7), 1:250 (dystrophin) or 1:400 (eMHC) (Jackson ImmunoResearch

Laboratories, Inc., West Grove, PA). Immunohistochemical methods were adapted from previously published methods from our lab (28, 45). Briefly, slides were dried and fixed in acetone for 10 min followed by several washes in 1X phosphate-buffered saline (PBS). Sections were then blocked with serum for 1 hour and incubated with 70 µg/ml goat anti-mouse monovalent Fab fragments (1:20, AffiniPure Fab Fragment Goat Anti-Mouse IgG (H+L),

Jackson ImmunoResearch Laboratories, Inc., West Grove, PA) for 30 min. Following blocking, sections were incubated in the primary antibody for overnight at 4°C (Pax7), 1 hour at room temperature (eMHC) or 30 min at 37°C (Dystrophin). After several washes with 1% BSA in

PBS, sections were then incubated in the appropriate secondary antibodies. Sections were washed with 1% BSA and stained with 4′,6-diamidino-2-phenylindole (DAPI, 1:20000) (Sigma-

Aldrich, St. Louis, MO) prior to mounting with Vectashield (Vector Laboratories, Burlingame,

CA).

Immunohistochemistry Analysis. Immunohistochemical images were obtained at 20X magnification for measurement of centrally located nuclei (CLN) and myofiber cross sectional area (CSA) and 40X for myonuclei to fiber ratio (N:F ratio) using a Leica DMRXA2 microscope. Images were acquired using a Zeiss AxioCam digital camera and Axiovision software (Zeiss, Thornwood, NY, USA). For assessing satellite cell content, slides were imaged and captured on the NanoZoomer digital slide scanner (Hamamatsu Corporation, Bridgewater,

NJ) using the 40X objective setting. Investigators were blinded to sample information for all assessments.

Immunohistochemistry Quantification. Dystrophin and DAPI stains were performed to quantify the CLN, myofiber CSA and N:F ratio (3 sections/sample). To assess CLN, total number of fibers, and the number of fibers with CLN were manually counted. At least 1,000 fibers per sample were counted. The results for each sample were then averaged. To assess myofiber CSA,

Adobe Photoshop was used to quantitate images acquired with a Ziess AxioCam digital camera.

Briefly, Dystrophin-FITC images acquired at 20X magnification from each sample were imported into Adobe Photoshop (CS5 Extended) where an average of 500 fibers per sample were manually circled using the magnetic lasso tool which grabs the positively stained pixels and decreases subjectivity and inter-assessment error. CSA for each fiber was recorded in the measurement log. The results for each sample were then averaged. To determine the myonuclei content, the number of nuclei for each myofiber was counted. Briefly, myofibers were manually counted by using Adobe Photoshop count tool. Total number of nuclei associated with each pre- numbered myofiber was then manually counted. To make the distinction between myonuclei and

54 mononuclear cells in the interstitial space, the nuclei that had a clear dystrophin line on only one side, and were contained within the myofiber were counted. 200 fibers per sample were analyzed.

The results for each sample were then averaged. Quiescent and activated satellite cells were detected with Pax7 antibody. The total number of Pax7+ cells per tissue section was manually counted. Only Pax7+ and DAPI+ cells associated with an individual myofiber were counted. Non- fiber associated nuclei (interstitial nuclei) were not included in the quantification. All data are represented as a percentage of Pax7+ cells to total myonuclei. The average number was assessed for three sections per animal.

Western Blot Analysis. All frozen gastrocnemius-soleus complexes collected after exercise training were manually ground with a mortar and pestle. Powdered tissue was homogenized in ice-cold buffer as previously described (45). Protein concentration was determined with the

Bradford protein assay using BSA for the standard curve. Equal amounts of protein (60 µg) were reduced, separated by SDS-PAGE using 6-8% acrylamide gels, and transferred to nitrocellulose membranes. Equal protein loading was verified by Ponceau S staining. Membranes were blocked in Tris-buffered saline (pH 7.8) containing 5% BSA and then were incubated with the appropriate phospho-specific antibody (phospho-mTOR, phospho-p70s6k) overnight at 4°C

(1:1000, Cell Signaling Technology, Danvers, MA). After multiple washes, HRP-conjugated anti-rabbit secondary antibodies (1:2000, Jackson ImmunoResearch, West Grove, PA) were applied to the blots for 1 hour at room temperature. Bands were detected using ECL western blotting substrate (SuperSignal West Dura; Thermo Fisher Scientific, Rockford) and a Bio-Rad

ChemiDoc XRS system (Bio-Rad, Hercules, CA). Bands were quantified using Quantity One software (Bio-Rad). After detection of phosphorylated proteins, blots were washed and stripped with Restore Western Blot Stripping Buffer (Thermo Fisher Scientific, Rockford, USA) for 20

55 minutes at room temperature. Membranes were washed and re-probed with total-specific antibodies (mTOR, p70s6k; 1:1000, Cell Signaling Technology, Danvers, USA) in the same manner as phospho-specific protein detection.

Hindlimb Grip Strength. Grip strength measurements for hindlimbs were tested once a week by using an electronic grip strength meter (Columbus Instruments, Columbus, OH). All mice were trained with the meter for three days prior to the initiation of the study. Each week, 24 hours following the last exercise session, hindlimb strength was measured. To assess strength, mice were passed over a metal mesh grid connected to a force transducer. Mice gripped the bar with their hindlimbs and then were tugged gently away until the grip was released. The peak force in grams was recorded by the transducer. Three trials were measured on each mouse during each test. The highest of the three values for each was recorded for analysis. Total body weight was measured before each grip strength test. The individual strength was indicated as limb strength

(g) divided by body weight (g). All measurements were performed by the same blinded examiner.

Statistical Analysis. All averaged data are presented as means ± SEM. To determine significance, comparisons between groups were performed by one-way ANOVA. The difference from baseline was used to determine significance for grip strength. When differences were observed,

Tukey’s post hoc analyses were performed. All calculations were with SPSS statistical software

(20.0, Chicago, IL). Differences were considered significant at P<0.05.

3.4 RESULTS

Sca-1+CD45- mMSCs Localize to Regenerating Myofibers Following Eccentric Exercise

The experimental design for examination of mMSC contribution to muscle repair and growth is provided in Figure 3.1A. The majority of fluorescently-labeled mMSCs were randomly present in muscle interstitium at two weeks post-injection. Consistent with observations previously reported at one week post-injection (42), we observed that mMSCs migrated to previously injured myofibers, identified by the presence of centrally located nuclei

(CLN) (Figure 3.1B). At two weeks, the amount of DiI-labeled cells in muscle tissue between mMSCs/SED and mMSCs/TR was not visibly different. mMSC Transplantation Allows for a Reduction in Body Weight During Four Weeks of Exercise

Training

While body weight and muscle weight were not significantly altered in any group at two weeks, body weight was lower in Saline/TR and mMSC/TR compared to Saline/SED at four weeks (P<0.05) (Table 3.1). The percent change in body weight at four weeks compared to baseline reduced in mMSC/TR compared to both sedentary groups (P<0.05) (Table 3.1). No significant group differences in absolute or relative muscle weight were observed following either two or four weeks of eccentric exercise training (Table 3.1). mMSC Transplantation Enhances Resolution of Muscle Damage at Two Weeks Post-Exercise

Training

All CLN+ fibers in the muscle were large, not small and triangular, at two and four weeks.

Therefore, we used this measure as an indicator of fibers resolving damage, rather than an indicator of a newly synthesized fiber. This was substantiated by our inability to detect embryonic myosin heavy chain positive fibers in any group at two or four weeks (Data not

57 shown). The percentage of myofibers with CLN was elevated 1.7 and 1.6-fold in Saline/TR compared to both Saline/SED and mMSC/SED respectively (P<0.05) (Figure 3.2A). Minimal

CLN+ fibers were detected in mMSC/TR (Fig 2A) at two weeks. Consistent with CLN data, macrophage content, as assessed by F4/80 immunostaining, was also significantly increased in

Saline/TR, but not in mMSC/TR compared to Saline/SED at two weeks (P<0.05) (Figure 3.2B).

The percentage of myofibers with CLN and macrophage content were no longer elevated in

Saline/TR or any group at four weeks (data not shown). mMSC Transplantation Significantly Enhances Satellite Cell Number Following Two Weeks of

Eccentric Exercise Training

We previously reported that satellite cell number was increased seven days after mMSC transplantation in muscle and this effect is further elevated if recipient mice are exercised prior to injection (42). In the current study, Pax7+ cell number was 1.7-fold higher in mMSC/TR compared to all other groups at two weeks (P<0.05) (Figure 3.3A). Satellite cells were quantified using Pax7/Dystrophin co-staining to ensure that all Pax7+ cells were in the satellite cell niche. Pax7+ cell number was no longer elevated in mMSC/TR or any group at the end of the four week training program (data not shown).

Myonuclear Content is Increased with mMSC Transplantation Following Eccentric Exercise

Training

Myonuclear accretion as a result of satellite cell fusion with existing myofibers can potentially contribute to muscle growth post-exercise training (7, 19). Consistent with our previously reported results, we did not observe direct fusion of DiI-labeled mMSCs with existing fibers (42). To assess overall myonuclear content, we counted the number of myonuclei per fiber

(N:F ratio). The N:F ratio was significantly higher in mMSCs/TR (2.08 ± 0.13) compared to all

58 other groups following two weeks exercise training (P<0.05) (Figure 3.3B). The N:F ratio was further elevated following four weeks of exercise training in mMSCs/TR (2.36 ± 0.12) compared to Saline/SED (P<0.05) (Figure 3.3C). No significant differences were detected in Saline/TR compared to other groups at two or four weeks (Figure 3.3C). mMSC Transplantation Increases Myofiber Hypertrophy and Hindlimb Strength Following

Eccentric Exercise Training

The mean fiber cross sectional area (CSA) was increased by 26% in mMSC/TR compared to Saline/TR following two weeks of downhill running exercise (P<0.05) (Figure

3.4A). Average myofiber CSA was also increased in mMSC/TR compared to Saline/SED (23%) and mMSCs/SED (17%) following four weeks of downhill running exercise (P<0.05) (Figure

3.4B). A rightward shift in the fiber size distribution was observed in mMSC/TR at four weeks, reflecting an increase in the percentage of large caliber fibers (Figure 3.4C). The percentage of fibers ranging >3000 µm2 was significantly increased in mMSC/TR compared to all the other groups (P<0.05) (Figure 3.4C).

Since gastrocnemius muscle has been shown to contribute to hindlimb grip strength (31), we used grip strength as a non-invasive measure of hindlimb function during the course of the four week study. This measurement allowed us to repeatedly and non-invasively assess force development as training progressed. Consistent with the increase in average fiber CSA, we observed a trend for increase in absolute hindlimb grip strength in mMSC/TR compared to all other groups (P=0.089) (Figure 3.5A). When normalized to body mass (g/g), we observed a 28% improvement in mMSC/TR compared to Saline/SED and mMSC/SED (P<0.05) (Figure 3.5B).

The percent change in grip strength after four weeks of exercise training compared to baseline

59 was also significantly higher in mMSCs/TR compared to both sedentary groups (P<0.05)

(Figure 3.5C).

Hypertrophic Signaling is Upregulated with mMSC Tranplantation Following Four Weeks of

Eccentric Exercise Training

mTOR-p70S6K signaling has been shown to play an integral role in the initiation of protein synthesis following mechanical stimuli (20, 22, 23). Consistent with small and insignificant increases in CSA and grip strength in Saline/TR at four weeks, mTOR and p70S6K phosphorylation (normalized to total protein) remained unchanged (Figure 3.6A and B).

However, both mTOR and p70S6K phosphorylation were increased almost 2-fold (P<0.05) in mMSCs/TR at four weeks compared to all other groups.

3.5 DISCUSSION

Our previous work has established that the α71 integrin is an important regulator of beneficial adaptations to eccentric exercise, including protection from damage, accelerated regeneration, and myofiber hypertrophy (5, 6, 28, 45). Recently, we reported that Sca-1+CD45- cells increase in skeletal muscle within 24 hours following an acute bout of eccentric exercise and this event was highly dependent on the presence of the 7 integrin (42). After characterizing the cells as multipotent MSCs, we completed short-term transplantation experiments to demonstrate that mMSCs derived from α7Tg muscle had the capacity to stimulate satellite cell proliferation, increase new fiber synthesis, and facilitate vascular growth in response to an acute bout of eccentric exercise (26, 44). In the current study, we sought to determine the extent to which mMSCs could regenerate skeletal muscle in response to multiple bouts of eccentric exercise. Here we demonstrate that a single injection of mMSCs isolated from α7Tg muscle can effectively enhance the rate of repair, myofiber growth and function following eccentric exercise training in adult WT mice.

Bone marrow-derived MSC migration to injured skeletal muscle is governed by a diverse set of cytokines and chemokines (18). The fact that human MSCs express a variety of chemokine receptors, including CCR1, CCR7, CCR9, CXCR4, CXCR5 and CXCR6, suggest that MSCs have the capacity to become mobilized and migrate to specific tissues based on unique chemokine gradients created following injury (21). Whether eccentric exercise preferentially stimulates tissue resident MSC proliferation or allows for recruitment of MSCs from distal tissues is not known. We have previously demonstrated that eccentric exercise can stimulate the proliferation of MSCs in muscle within 24 hours (24), yet recruitment from bone marrow or another tissue source cannot be ruled out. Despite this gap in knowledge, we consistently observe

61 localization of transplanted mMSCs in close proximity to large CLN+ fibers at one and two weeks following exercise ((42) (Figure 3.1A)). This observation would suggest the existence of a chemical gradient necessary for mMSC migration to the interstitium of damaged fibers following exercise. However, the fact that eccentric exercise can stimulate MSC accumulation in skeletal muscle which is resistant to damage (MCK: α7BX2 integrin) suggests that other factors, including myofiber strain or force development, may similarly allow for the release of cytokines or chemokines that can positively influence MSC recruitment, migration, proliferation, and/or stromal function. In vitro experiments utilizing transwell assays to evaluate the MSC migratory response to conditioned media from damaged versus stretched fibers may provide some light on exercise-specific microenvironmental cues.

Transplantation of bone marrow-derived MSCs can effectively repair muscle and restore skeletal muscle function in response to a variety of different types of injury, including resection, crush, or chemical toxin-based injuries (11, 17, 33, 43, 44). In this study, our goal was not to determine the extent to which bone marrow-derived MSCs could repair damage in response to exercise, but rather assess endogenous MSC contribution to regeneration to account for differences in MSC function between tissue types, including the potential for MSCs to directly become myogenic. Consistent with previous observations, we did not detect fusion of dye- labeled MSCs with existing fibers, yet a single injection of a limited number of cells

(40,000/muscle) into a relatively large muscle (gastrocnemius), resulted in significant improvements in resolution of damage within two weeks. mMSCs were still present in the muscle at two weeks post-injection, suggesting long-term capacity for healing.

Although the lack of large CLN+ fibers and macrophage presence with transplantation suggests that damage was indeed resolved at two weeks following the initiation of exercise, it is

62 possible that mMSC transplantation minimized the onset of damage, including secondary injury associated with inflammation. Therefore, although the lack of damage and inflammation observed in 7Tg muscle post-exercise is assumed to occur as a result of increased adhesion between the extracellular matrix and the cytoskeleton within areas that are particularly mechanosensitive (Z bands, myotendinous junction), we cannot exclude the possibility that mMSC accumulation and perhaps MSC secretion of anti-inflammatory cytokines may directly account for these positive adaptations. Short-term (24 hour) evaluations of MSC influence on exercise-induced changes in sarcolemmal integrity, ultrastructural damage, immune response and functional deficits are necessary to address this important question.

In our previous study, we found that mMSC injection 24 hours following a single bout of eccentric exercise significantly increased satellite cell content in skeletal muscle, whereas mMSC supplementation in sedentary mice had no effect. In the current study, we similarly demonstrate enhancement of satellite cells with mMSC injection only in combination with multiple bouts of eccentric exercise (Figure 3.3A). Therefore, MSCs alone are not capable of stimulating resident progenitors in healthy muscle; rather injury or other factors associated with eccentric exercise are a requirement for MSC activation and contribution to Pax7+ cell expansion. The rise in Pax7+ cell number at two weeks post-exercise was unexpected given the absence of damage and inflammation in mMSC injected muscle. The prolonged elevation of satellite cell content in the absence of damage and inflammation is consistent with previous observations in 7Tg mice (28).

These data imply that while the innate immune system, including alternative macrophages, provides important cues to activate satellite cells, MSCs also support satellite cell proliferation, differentiation, and/or survival.

mMSC transplantation elicited an increase in myofiber growth four weeks following initiation of exercise training. Although the changes observed were small, the increase in fiber

CSA was consistent with physiological growth observed in rodents following wheel running with resistance (~20-30%) (Figure 3.4A-D) (27). We hypothesized that satellite cell expansion at two weeks post-transplantation may increase the opportunity for myoblast fusion. The nuclei:fiber ratio (N:F) was elevated as early as two weeks post-exercise in mMSC-treated muscle and the ratio was further increased at four weeks (Figure 3.3D&E). As previously stated, we did not observe fusion of DiI+ cells with fibers, either in the cytoplasm or localized to nuclei at any time points in the current study. Thus, myoblast fusion likely accounted for myofiber growth observed in this group. Although we can only speculate based on the fact that signaling was analyzed on whole tissue samples, the 2-fold increases in both mTOR and p70s6k phosphorylation may reflect an increase in protein synthesis associated with enhanced repair, enriched myonuclear content, and/or increased translational capacity associated with MSC paracrine factor release.

Regardless of the precise mechanism responsible for growth, these changes may have contributed to the significant improvement in relative tension observed in MSC injected mice at four weeks post-exercise (Figure 3.5B).

We previously evaluated our Sca-1+CD45- mMSCs for satellite cell markers as well as a variety of non-satellite cell markers via flow cytometry and PCR (42). While sorting non- satellite cells based on two markers, Sca-1 (positive selection) and CD45 (negative selection), increases the potential for retrieving a heterogenous mononuclear cell population, we found that mMSCs do not express Pax7, yet are positive for several MSC (CD90, CD105, CD29, CD73) and pericyte markers (NG2, PDGFR, CD146). Although not reported in our previously published findings, mMSCs are also positive for PDGFR. The extent to which these cells

64 represent type 1 pericytes (Sca-1+NG2+PDGFRα+) (4), FAPs (Sca-1+PDGFR+lin-) (25), or PICs

(Sca-1+PW1+PDGFRα+) (34) previously reported in the literature is not known, yet likely provided their limited capacity for myogenesis and stromal contribution to repair (42). MSCs, true to their mesodermal origin, have the capacity to differentiate into adipocytes and fibroblasts.

Since mechanical strain can inhibit MSC adipogenesis (40), it will be important to determine the extent to which tension created in the muscle during eccentric exercise has the capacity to promote multipotency while maintaining a secretome that supports tissue health.

In conclusion, this study provides the first evidence of a role for mMSCs in long-term skeletal muscle repair and growth following repeated bouts of eccentric exercise. Further studies are necessary to determine the origin of MSCs that accumulate in muscle post-exercise, the factors in the microenvironment that regulate mMSC migration and function, and the precise mechanisms by which mMSCs contribute to the adaptive response to eccentric exercise. We believe this information will guide us towards a better understanding of the basis for skeletal muscle atrophy and development of interventions that can effectively preserve muscle mass and function with age, immobilization and disease.

3.6 ACKNOWLEDGEMENTS

We would like to thank Anthony Zhang and Zak Kammer for their assistance in data analysis.

This work was supported by a grant from the Ellison Medical Foundation (to MDB). HDH was supported by a National Science Foundation Integrative Graduate Education and Research

Traineeship (IGERT) in Cellular and Molecular Mechanics and BioNanotechnology (CMMB).

None of the authors declared any conflict of interest.

3.7 FIGURES, TABLE AND FIGURE LEGENDS

Figure 3.1. Sca-1+CD45- mMSCs localize to regenerating myofibers following eccentric exercise training. (A) Experimental design for injection of saline or DiI-labeled mMSCs into gastrocnemius muscle followed by 2 or 4 weeks of exercise training. (B) Representative cross-section of a gastrocnemius muscle from skeletal muscle transplanted with mMSCs at 2 weeks post-exercise. Arrows indicate DiI- labeled mMSCs at high resolution. Expression of DiI (Red), Dystrophin (Green), DAPI (Blue) and merged images at 20X (scale bar = 20 m) and 40X (scale bar = 40 m) magnification are illustrated.

Table 3.1. Body and Muscle Weight. For two weeks study, saline was injected to contralateral leg of the same animal used for mMSC transplantation. BW, body weight; BW%, percent change of body weight; MW, muscle weight, rMW, relative muscle weight (normalized to body weight). * P < 0.05 compared with Saline/SED; § P < 0.05 compared with mMSC/SED. n = 5-6/group. Values are presented as mean ± SEM.

Figure 3.2. mMSC transplantation enhances resolution of muscle damage following eccentric exercise training. (A) Quantitation and visualization of myofibers with centrally-located nuclei (CLN) following saline or mMSC injection at two weeks; sedentary (SED) or exercise training (TR). FITC (green) = dystrophin, DAPI (blue) = nuclei (arrows), 20X, scale bar = 20 m. (B) Quantitation and visualization of F4/80 positive cells following saline or mMSC injection. FITC (green) = F4/80, DAPI (blue) = nuclei (arrows), 20X, scale bar = 20 m. * P < 0.05 compared with Saline/SED; § P < 0.05 compared with mMSC/SED. n = 5-6/group. Values are presented as mean ± SEM.

Figure 3.3. mMSC transplantation significantly enhances satellite cell number and myonuclear content following eccentric exercise training. (A) Quantitation and visualization of Pax7+ satellite cells following saline or mMSC injection at two weeks. FITC (green) = Dystrophin, TRITC (Red) = Pax7 (arrows), 40X, scale bar = 10 m. Quantitation of the number of myonuclei per fiber following saline or mMSC injection at (B) two and (C) four weeks. * P < 0.05 compared with Saline/SED; # P < 0.05 compared with Saline/TR; § P < 0.05 compared with mMSCs/SED. n = 5-6/group. Values are presented as mean ± SEM.

Figure 3.4. mMSC transplantation increases myofiber hypertrophy following eccentric exercise training. Mean cross-sectional area (CSA) of muscle fibers in gastrocnemius muscle following saline or mMSC injection at (A) two or (B) four weeks. (C) Muscle fiber CSA distribution at four weeks. * P < 0.05 compared with Saline/SED; # P < 0.05 compared with Saline/TR; # P < 0.05 compared with Saline/TR; § P < 0.05 compared with mMSCs/SED. n = 5-6/group. Values are presented as mean ± SEM.

Figure 3.5. mMSC transplantation increases hindlimb strength following four weeks of eccentric exercise training. Hindlimb grip strength was assessed every week. (A) Absolute hindlimb grip strength (peak tension, g). (B) Relative hindlimb grip strength (peak tension/body weight, g/g). (C) Percentage changes in relative hindlimb grip strength after four weeks training compared to sedentary state. * P < 0.05 compared with Saline/SED; § P < 0.05 compared with mMSCs/SED. n = 5-6/group. Values are presented as mean ± SEM.

Figure 3.6. Hypertrophic signaling is upregulated with mMSC transplantation following four weeks of eccentric exercise training. (A) Representative blot and densitometric analysis of mTOR phosphorylation (Ser2448) and protein expression. S = saline, M = mMSC, SED = sedentary, TR = four weeks of exercise training. (B) Representative blot and densitometric analysis of p70S6K phosphorylation (Thr389) and protein expression. Fold changes relative to Saline/SED are presented. * P < 0.05 compared with Saline/SED; # P < 0.05 compared with Saline/TR; § P < 0.05 compared with mMSCs/SED. n = 5- 6/group. Values are presented as mean ± SEM.

3.8 REFERENCES

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26. Kadi F, Charifi N, Denis C, Lexell J, Andersen JL, Schjerling P, Olsen S, and Kjaer M. The behaviour of satellite cells in response to exercise: what have we learned from human studies? Pflugers Arch 451: 319-327, 2005. 27. Konhilas JP, Widegren U, Allen DL, Paul AC, Cleary A, and Leinwand LA. Loaded wheel running and muscle adaptation in the mouse. Am J Physiol Heart Circ Physiol 289: H455- 465, 2005. 28. Lueders TN, Zou K, Huntsman HD, Meador B, Mahmassani Z, Abel M, Valero MC, Huey KA, and Boppart MD. The alpha7beta1-integrin accelerates fiber hypertrophy and myogenesis following a single bout of eccentric exercise. Am J Physiol Cell Physiol 301: C938- 946, 2011. 29. Malm C, Sjodin TL, Sjoberg B, Lenkei R, Renstrom P, Lundberg IE, and Ekblom B. Leukocytes, cytokines, growth factors and hormones in human skeletal muscle and blood after uphill or downhill running. J Physiol 556: 983-1000, 2004. 30. Maughan RJ, Donnelly AE, Gleeson M, Whiting PH, Walker KA, and Clough PJ. Delayed-onset muscle damage and lipid peroxidation in man after a downhill run. Muscle Nerve 12: 332-336, 1989. 31. Maurissen JP, Marable BR, Andrus AK, and Stebbins KE. Factors affecting grip strength testing. Neurotoxicol Teratol 25: 543-553, 2003. 32. McCarthy JJ, Mula J, Miyazaki M, Erfani R, Garrison K, Farooqui AB, Srikuea R, Lawson BA, Grimes B, Keller C, Van Zant G, Campbell KS, Esser KA, Dupont- Versteegden EE, and Peterson CA. Effective fiber hypertrophy in satellite cell-depleted skeletal muscle. Development 138: 3657-3666, 2011. 33. Merritt EK, Cannon MV, Hammers DW, Le LN, Gokhale R, Sarathy A, Song TJ, Tierney MT, Suggs LJ, Walters TJ, and Farrar RP. Repair of traumatic skeletal muscle injury with bone-marrow-derived mesenchymal stem cells seeded on extracellular matrix. Tissue Eng Part A 16: 2871-2881, 2010. 34. Mitchell KJ, Pannerec A, Cadot B, Parlakian A, Besson V, Gomes ER, Marazzi G, and Sassoon DA. Identification and characterization of a non-satellite cell muscle resident progenitor during postnatal development. Nat Cell Biol 12: 257-U256, 2010. 35. Motohashi N, Uezumi A, Yada E, Fukada S, Fukushima K, Imaizumi K, Miyagoe- Suzuki Y, and Takeda S. Muscle CD31(-) CD45(-) side population cells promote muscle regeneration by stimulating proliferation and migration of myoblasts. Am J Pathol 173: 781-791, 2008. 36. Nosaka K, and Newton M. Repeated eccentric exercise bouts do not exacerbate muscle damage and repair. J Strength Cond Res 16: 117-122, 2002. 37. Relaix F, and Zammit PS. Satellite cells are essential for skeletal muscle regeneration: the cell on the edge returns centre stage. Development 139: 2845-2856, 2012.

38. Sambasivan R, Yao R, Kissenpfennig A, Van Wittenberghe L, Paldi A, Gayraud- Morel B, Guenou H, Malissen B, Tajbakhsh S, and Galy A. Pax7-expressing satellite cells are indispensable for adult skeletal muscle regeneration. Development 138: 3647-3656, 2011. 39. Schoenfeld BJ. Does exercise-induced muscle damage play a role in skeletal muscle hypertrophy? J Strength Cond Res 26: 1441-1453, 2012. 40. Sen B, Xie Z, Case N, Ma M, Rubin C, and Rubin J. Mechanical Strain Prevents Adipogenesis in Mesenchymal Stem Cells By Stimulating a Durable beta-Catenin Signal. J Bone Miner Res 23: S232-S233, 2008. 41. Uezumi A, Fukada S, Yamamoto N, Takeda S, and Tsuchida K. Mesenchymal progenitors distinct from satellite cells contribute to ectopic fat cell formation in skeletal muscle. Nat Cell Biol 12: 143-152, 2010. 42. Valero MC, Huntsman HD, Liu J, Zou K, and Boppart MD. Eccentric exercise facilitates mesenchymal stem cell appearance in skeletal muscle. PLoS One 7: e29760, 2012. 43. von Roth P, Duda GN, Radojewski P, Preininger B, Perka C, and Winkler T. Mesenchymal stem cell therapy following muscle trauma leads to improved muscular regeneration in both male and female rats. Gend Med 9: 129-136, 2012. 44. Winkler T, von Roth P, Radojewski P, Urbanski A, Hahn S, Preininger B, Duda GN, and Perka C. Immediate and delayed transplantation of mesenchymal stem cells improve muscle force after skeletal muscle injury in rats. J Tissue Eng Regen Med 6 Suppl 3: s60-67, 2012. 45. Zou K, Meador BM, Johnson B, Huntsman HD, Mahmassani Z, Valero MC, Huey KA, and Boppart MD. The alpha(7)beta(1)-integrin increases muscle hypertrophy following multiple bouts of eccentric exercise. J Appl Physiol (1985) 111: 1134-1141, 2011.

CHAPTER 4

LAMININ-111 ENHANCES THE MYOGENIC RESPONSE TO ECCENTRIC

EXERCISE IN YOUNG AND AGED MICE

4.1 ABSTRACT

Aging is associated with a loss of regenerative capacity in muscle that contributes to the onset and progression of sarcopenia. Alterations in the muscle microenvironment are at least partially responsible for these impairments in muscle regeneration with age. Laminin-111 (LM-

111), an important component of the extracellular matrix (ECM) during embryonic development, has been demonstrated to enhance myogenesis in mouse models of muscular dystrophy; however, the effects of LM-111 supplementation in young healthy and aged muscle have not been investigated. The purpose of the present study was to determine if LM-111 supplementation could enhance muscle repair following eccentric exercise in both young and aged muscle. LM-

111 or Saline (SA) was injected intramuscularly into either 3-month-old (Young) or 23-month- old (Aged) C57Bl/6 mice. Mice either remained sedentary (SED) or were subjected to a single bout of downhill running (EX) to induce muscle damage. In young mice, satellite cell number was significantly higher in LM-111/EX compared all other groups (P<0.05), primarily due to enhanced satellite cell proliferation (P<0.05). LM-111 increased myoblast quantity as compared to collagen when treated in culture (P<0.05). LM-111 injection also increased the quantity of eMHC+ fibers in young mice post-eccentric exercise (P<0.05). In aged mice, LM-111 injection prevented muscle force loss compared to saline-injected controls (P<0.05); however, satellite cell quantity only trended to increase (P=0.09). The enhanced myogenic response to LM-111 injection may in part be due to altered genetic expression of a variety of paracrine factors in

78 muscle-derived mesenchymal stem cells (mMSCs) in both young and old mice. Together, these data demonstrate that LM-111 supplementation enhances the regenerative response to eccentric muscle damage in both young and aged muscle.

Keywords: aging, α7 integrin, satellite cells, mesenchymal stem cell, downhill running, repair,

4.2 INTRODUCTION

Muscle aging is characterized by the loss of muscle mass and function, which is partially attributable to increased susceptibility to damage and impaired regeneration (10). Muscle regeneration is dependent on a functioning population of myogenic stem cells known as satellite cells. Satellite cells reside in a state of quiescence on the periphery of myofibers between the sarcolemma and basal lamina (12). In response to muscle damage, satellite cells become activated, proliferate and fuse to existing myofibers to facilitate muscle repair (24). Recently, a variety of non-satellite stem cells, expressing mesenchymal stem cells (MSCs) markers, have been identified in skeletal muscle that may contribute to muscle repair following injury via direct or indirect mechanisms (2, 27, 41, 43, 57). With advanced age, the satellite cell population becomes dysfunctional resulting in impaired muscle regeneration following damage and decreased hypertrophic response when exposed to anabolic stimuli (15). A combination of these factors has been implicated as underlying causes of sarcopenia.

Recent evidence suggests that alterations in the muscle microenvironment with age are responsible for the impaired regenerative response (6, 14). The extracellular matrix (ECM) is an important structural component for muscle fiber development and muscle tissue integrity, and provides a niche for muscle progenitor cells, and regulates the release of paracrine factors involved in the process of muscle repair (39). Laminins are biologically active ECM proteins found in the basal lamina of skeletal muscle fibers that provide an important scaffold necessary for muscle tissue organization, maintenance, and survival (29). Laminins are hetetotrimers formed by one heavy chain (α) and two light chains (β and γ) that combine in different ways to form fourteen isoforms (3). Among them, Laminin-111 (α1, β1, γ1) (LM-111) is the predominant isoform during embryonic skeletal muscle development but is no longer expressed

80 in adult skeletal muscle (23). The relationship between laminins and their receptors is thought to play a critical role for skeletal muscle function and integrity (62). The role of LM-111 in myogenesis has been extensively studied and found to be involved in primary myoblast proliferation, motility and myotube formation (21, 54), which has led to their use as therapy in pre-clinical models of muscular dystrophy (21, 49-51, 60). In the majority of these studies, LM-

111 injection has been demonstrated to inhibit muscle damage and enhance muscle regeneration through increased satellite cell expansion and new fiber (21, 49-51, 60). To date, only a single study has failed to show a beneficial response to laminin therapy (19). In the study by Gawlik and colleagues, only laminin α1 was genetically overexpressed, which differs from other studies that injected full-length LM-111 (21, 49-51, 60) . Although the effects of LM-111 therapy have been well-established in dystrophic mouse models, the effect of LM-111 in non-dystrophic muscle tissue has not been studied, neither in the response to exercise-induced muscle damage or aging.

LM-111 is a preferred ligand for α7β1 integrin (44), and an increase in α7 integrin levels has been suggested as one of the key mechanisms responsible for restoring muscle regeneration

(49, 50, 60). Recently we have demonstrated that transgenic overexpression of the α71 integrin in mouse muscle suppressed macrophage infiltration, sarcolemmal damage, increases in new fiber synthesis, and satellite cell number following single bout of downhill running (35, 63).

Furthermore, we established that Sca-1+CD45- MSCs are increased in muscle (mMSCs) following eccentric exercise in an α7 integrin-dependent manner (59), and that these mMSCs contribute to muscle regeneration following eccentric exercise after transplanting into wild-type mouse muscles (59). These data suggest that the accumulation of mMSCs may be a significant contributing factor to the enhanced muscle regeneration observed in α71 transgenic mice.

Additionally, we have recently demonstrated that mMSCs alter their cell surface phenotype when cultured on laminin compared to gelatin (59), and that culture on laminin promotes the release of factors from mMSCs that enhance vasculogenesis (25) and myogenesis (De Lisio et al. in prep). Together, these data suggest that by increasing levels of α71 integrin, LM-111 may enhance muscle repair after a damaging bout of eccentric downhill running exercise (13).

Whether LM-111 injection can increase the expression of α7 integrin in healthy young or old muscle tissue similar to the transgenic model is unknown, but could lead to the therapeutic use of

LM-111 to inhibit sarcopenia.

The purpose of the present investigation was to determine if LM-111 therapy could enhance the acute myogenic response to eccentric muscle damage induced by downhill running in both young and old mice. Given the role of LM-111 in myogenesis and its successful application in pre-clinical models of muscular dystrophy, we hypothesized that LM-111 therapy would increase satellite cell quantity immediately post-eccentric exercise in both young and old mice, and that the mechanism responsible for this response would be due, in part, to increased production of paracrine factors from mMSCs.

4.3 METHODS

Animals. Protocols for animal use were approved by the Institutional Animal Care and Use

Committee (IACUC) of the University of Illinois at Urbana-Champaign. Three-month-old

(young) female (Charles River) and 23-month-old (old) male C57Bl/6 mice were purchased from the National Institute of Aging (NIA). All mice were housed in a temperature-controlled animal room, maintained on a 12:12 light-dark and provided standard laboratory chow and water ad libitum.

Laminin-111 Injection. Natural mouse Laminin-111 (α1, β1, γ1; LM-111) purified from

Engelbreth-Holm-Swarm mouse sarcoma cells (Invitrogen Life Technologies, Grand Island, NY,

USA) was thawed overnight at 4°C. At Day 0, each mouse was injected intramuscularly (i.m.) into both gastrocnemius (GAS) muscles with either 100 µL 1mg/ml EHS LM-111 (LM-111, n=12) or 100 µL sterile TBS (50 mM Tris-HCL (pH 7.4), 0.15 M NaCl in PBS) (SA, n=12)

(Figure 1A & 6A) (51). Two separate injections, each of 50 µL, were injected into the lateral and medial lobe of the gastrocnemius to ensure uniform distribution throughout the entire muscle.

Exercise Protocol and Tissue Harvest. At either 1 week (young) (Figure 4.1A) or 3.5 weeks (old)

(Figure 4.6A) post-injection, half of the mice in each group (saline or LM-111 injected) underwent a single 30 minute bout of downhill running exercise (EX, n=6/group) to induce muscle damage on a motorized treadmill (Exer-6M, Columbus Instruments, Columbus, OH) or remained sedentary (SED, n=6/group). Exercise was conducted as previously described (5, 34).

Briefly, mice ran at a 20o decline for 30 minutes at a max speed of 17 m/min for young mice or

13-15 m/min for old mice. Each session consisted of a warm-up period where speed was gradually increased by 1m/min from 10 m/min to the final target speed. SED mice were place on the treadmill for 30 minutes to control for the stress of exercise. Old mice were not exercised

83 until 3.5 weeks post-injections to allow for maximal improvement of the skeletal muscle environment from LM-111 injections, and LM-111 has been demonstrated to be present in skeletal muscle up to 28 days following a single injection (49, 50, 60). Twenty-four hours following the eccentric exercise, all mice were euthanized via carbon dioxide asphyxiation and both gastrocnemius-soleus complexes were rapidly dissected. One gastrocnemius-soleus complex was used for fluorescence-activated cell sorting (FACS) to isolate Sca-1+CD45- mMSCs and the other was cut in half and either frozen in liquid nitrogen for protein analysis or pre-cooled isopentane for immunohistochemical analysis.

Isolation of Sca-1+CD45- Cells from Skeletal Muscle. Sca-1+CD45- cells were isolated via

FACS, as previously described (59). Briefly, the gastrocnemius-soleus complex was resected, minced and enzymatically digested. Filtered samples were incubated on ice with anti-mouse

CD16/CD32 (1 μg/106 cells) (eBioscience, San Diego, CA) for 10 min in order to block non- specific Fc-mediated interactions. Cells were then stained in a cocktail of monoclonal anti-mouse antibodies: Sca-1-PE (600ng/106 cells) and CD45-APC (300 ng/106 cells) (eBioscience, San

Diego, CA) diluted in 2% FBS in PBS. FACS was performed for Sca-1+CD45- cell quantification and isolation using an iCyt Reflection System (Urbana, IL). Negative and single- stained controls were used to establish gates for antibody staining and low forward and back scatter events were excluded as debris. Sorted cell suspensions were immediately centrifuged, resuspended in of RLT buffer (RNeasy Micro Kit) and frozen at -80oC until further analysis.

RNA extraction and RT-PCR. Total RNA was extracted from frozen sorted cells using the

RNeasy Micro Kit and QIAshredder spin columns according to the manufacturer's instructions

(Qiagen, Valencia, CA). The RNA concentrations and purities were analyzed spectrophotometrically by Synerg H1 Hybrid Multi-Mode Microplate Reader (Biotek, Winooski,

VT). The reverse transcriptase reaction was performed using first strand cDNA synthesis kit

(Applied Biosystems, Grand Island, NY) according to manufacturer’s instructions in thermo cycler (GeneAmp PCR Machine 9700, Applied Biosystems, Grand Island, NY). For detection of certain genes, cDNA was preamplified using TaqMan PreAmp Master Mix (Life Technologies

Inc, Grand Island, NY) according to manufacturer’s instructions. Briefly, all stock primer/probe mixes were prediluted together in Tris–EDTA buffer to give 0.2x concentration. The preamplification reagent comprised 25 µL TaqMan PreAmp Master Mix and 12.5 µL pooled primers mix to 0.2x concentration. After adding 12.5 µL cDNA, the tubes were placed in a PCR thermal cycler and incubated at 95°C for 10 min. Samples were preamplified for 14 cycles. The preamplified samples were then diluted 1:20 with Tris–EDTA buffer. cDNA was stored at -20oC until qPCR analysis. Gene expression was individually quantified for all genes of interest using a qRT-PCR machine (7900HT Fast Real-Time PCR system and SDS Enterprise Database) and analyzed using the CT method (Table 4.1). Expression of GAPDH was used as the housekeeping gene.

Immunohistochemistry. Muscle complexes from a single limb were divided at the midline along the axial plane, and the distal end was embedded in OCT (Tissue-Tek; Fischer Scientific) for immunohistochemistry analysis. Transverse cryosections (8μm) were cut for each histological assessment using a CM3050S cryostat (Lecia, Wezlar, Germany). Sections were placed on frozen microscope slides (Superfrost; Fischer Scientific, Hanover Park, IL) and stored at -80°C before staining. Sections were stained with antibodies against Laminin 1 (1:250, Millipore,

Billerica, MA); dystrophin (1:100, MANDRA-1, Sigma, St. Louis, MO); Pax7 (1:2,

Developmental Studies Hybridoma Bank, Iowa City, Iowa); eMHC (BF-45, 1:8, Developmental

Studies Hybridoma Bank, Iowa City, Iowa) and Ki67(1:100, Abcam, Cambridge, MA). All

85 species-appropriate secondary antibodies used for immunofluorescence studies were applied at

1:100 (Ki67), 1:200 (Laminin α1 and Pax7), 1:250 (Dystrophin) or 1:400 (eMHC) (Jackson

ImmunoResearch Laboratories, Inc., West Grove, PA). Immunohistochemical methods were adapted from previously published methods from our lab (35, 63). Briefly, slides were thawed and fixed in acetone for 10 minutes at -20oC followed by several washes in 1x phosphate- buffered saline (PBS). Sections were then permeabilized in 0.1% Triton-X/PBS and blocked in

PBS containing 5% BSA/5% goat anti-mouse monovalent Fab fragments /2% Goat Serum for 1 hour at room temperature or for 20 min or with blocking solution (10% FBS, 10% GS and 2.5%

BSA in PBS) for 60 min (Laminin 1) followed by 70 µg/ml goat anti-mouse monovalent Fab fragments (1:20, AffiniPure Fab Fragment Goat Anti-Mouse IgG (H+L), Jackson

ImmunoResearch Laboratories, Inc., West Grove, PA) for 30 min. Following blocking, sections were incubated in the primary antibody for either 30 min at 37°C (Dystrophin), 1 hour at room temperature (eMHC) or overnight at 4°C (Pax7). After several washes with 1% BSA in PBS, sections were then incubated in the appropriate secondary antibodies. Sections were then washed with 1% BSA and stained with 4′,6-diamidino-2-phenylindole (DAPI, 1:20000) (Sigma-Aldrich,

St. Louis, MO) for nuclear staining. For co-immunofluorescent staining, sections were re-fixed in 4% PFA (Sigma-Aldrich, St. Louis, MO) to prevent migration of the secondary antibodies and re-blocked in 10% GS in 0.01% Triton-X 100 prior to incubation with primary and secondary antibodies as described above.

Immunohistochemistry Analysis. Immunohistochemical images were obtained at either 20X or

40X-magnification using a Leica DMRXA2 microscope. Images were acquired with a Zeiss

AxioCam digital camera and Axiovision software (Zeiss, Thornwood, NY, USA). For assessing satellite cell content, slides were imaged and captured on the NanoZoomer digital slide scanner

(Hamamatsu Corporation, Bridgewater, NJ) using the 40X objective setting. Investigators were blinded to sample information for all assessments.

Immunohistochemistry Quantification. Pax7+ cells were quantified from an average of 896 myonuclei per sample by manual counting in ImageJ and data are presented as the percentage of

Pax7+ cells per myonuclei. Ki67+/Pax7+ cells were only counted as positive if they were triple immunolabeled with DAPI and antibodies against Pax7 and Ki67. Ki67+/Pax7+ cells are expressed as a percentage of total myonuclei, and the average number was assessed for three sections per animal.

Western Blot Analysis. The remaining proximal end of each muscle used for immunohistochemical analysis was manually ground with a mortar and pestle for isolation of proteins for western blot analysis. Powdered tissue was homogenized in ice-cold buffer (50 mMTris-HCl pH7.4, 150 mM NaCl, 1 mM PMSF, 1% NP-40, 0.5% Sodium deocycholate, 0.1%

SDS) containing phosphatase and protease inhibitors (Roche) as previously described (63).

Protein concentration was determined with the Bradford protein assay using BSA for the standard curve. Equal amounts of protein (30 µg) were separated by SDS-PAGE using 6-8% acrylamide gels, and transferred to nitrocellulose membranes under non-reducing conditions.

Equal protein loading was verified by Ponceau S staining. Membranes were blocked in Tris- buffered saline (pH 7.8) containing 5% non-fat dry milk overnight at 4 °C and then were incubated with the appropriate antibody for 1 hour at room temperature (1:1000, α7A: CDB345 and α7B: CDB347). After multiple washes, HRP-conjugated anti-rabbit secondary antibodies

(1:10000, Jackson ImmunoResearch, West Grove, PA) were applied to the blots for 1 hour at room temperature. Bands were detected using ECL western blotting substrate (SuperSignal West

Dura; Thermo Fisher Scientific, Rockford, Illinois) and a Bio-Rad ChemiDoc XRS system (Bio-

Rad, Hercules, CA). Bands were quantified using Quantity One software (Bio-Rad Hercules,

CA). Expression for each protein of interest was normalized to the expression of the housekeeping protein β-actin, which was blotted in a separate membrane.

Handgrip Strength Test. Grip strength measurements for all four limbs were tested weekly using an electronic Grip Strength Meter (Columbus Instruments, Columbus, OH). All mice were trained with the meter for three days prior to the initiation of the study. To assess strength, mice were passed over a metal mesh grid connected to a force transducer. Mice gripped the bar with both their fore- and hind-limbs and then were tugged gently away until they released their grip.

The peak force in grams was recorded by the transducer. Three trials were measured on each mouse during each test. Both peak and average force of the three trials were used for analysis.

Total body weight was measured before each grip strength test for calculation of relative strength. A blinded examiner conducted all force measurements for old mice.

In Vitro Primary myoblast Proliferation. Polydimethylsiloxane (PDMS) gel was used to coat plates (BioFlex plates, Flexcell International, McKeesport, PA, USA). Briefly, PDMS gel was prepared by mixing the elastomer base and curing agent (Sylgard 184, Dow Corning, Midland,

MI). The stiffness of the gel can be adjusted by adding curing agent in different ratios to the base.

In the present study, we used the curing agent in 1:50 (10kpa) to the base to mimic the stiffness of young muscle (32). The mixed PDMS gel was then added onto each well of flexcell plate (1ml) to cover the entire surface. The plates were placed in the oven for 3 hours at 65 to cure the gel.

Collagen I (BD Science, San Jose, CA) or LM-111 solution was added on top of PDMS gel and incubate 60 min at room temperature. Since cells do not directly attach to PDMS gel, same amount of total ECM protein was used to coat the plates for all conditions. Thus, to analyze the

LM-111 dose effect on primary myoblast proliferation, the following four conditions were used

88 in this study: 50 μg/ml Collagen I (control), 10 μg/ml LM-111 + 40 μg/ml Collagen I, 30 μg/ml

LM-111 + 20 μg/ml Collagen I, and 50 μg/ml LM-111. All wells were then incubated in 2mM glycine overnight at 4 to quench excess Aldehyde groups.

Myoblasts (5 x 104) were seeded onto LM-111 or collagen-coated plates in growth media

(DMEM supplemented with 10% fetal bovine serum, 5ng/ml recombinant human

FGFV(Invitrogen Life Technologies, Grand Island, NY), 100 U/ml penicillin, 100 μg/ml streptomycin) at 37°C with 5% CO2 for 96 hours. Upon 60-70% confluence, cells were incubated in serum-free DMEM for 18 hours. Half of the plates were then subjected to 15% cyclic strain for 1 hour at 1 Hz. 24 hours following strain, all cells were rinsed in PBS and trypsinized for proliferation assay analyzed by cell counting using a hemocytometer. The number presented is the average of six wells.

Statistical Analysis. All data are presented as means ± SEM. Comparisons between groups were performed by two-factor ANOVA (injection: saline or LM-111 and intervention: sedentary or exercise) followed by Tukey post-hoc test when significant differences were detected. One-way

ANOVA with Tukey post-hoc test was used to test significance of the in vitro cell proliferation experiments. A two-factor repeated-measures ANOVA was used to test for differences in body weight and grip strength in aged mice. All calculations were with either SPSS statistical software

(20.0, Chicago, IL) or SigmaPlot 12.5 (SYSTAT, Chicago, IL). Differences were considered significant at P<0.05.

4.4 RESULTS

LM-111 Improves the Regenerative Response to Eccentric Exercise in Young Skeletal Muscle

LM-111 was present throughout the muscles injected intramuscularly with LM-111, but was not observed in muscles injected with saline (Figure 4.1B). In LM-111 injected muscles,

LM-111 was primarily found outlining myofibers consistent with its role as a primary protein component of the basal lamina (Figure 4.1B). Furthermore, LM-111 appeared to co-localize with satellite cells as indicated by co-labeling with Pax7 (Figure 4.1C).

In young mice, satellite cell content, as indicated by Pax7 staining, was significantly increased in LM-111 injected mice compared to saline injected controls (LM-111 main effect,

P<0.05). In addition, satellite cell content was significantly increased in LM-111 injected mice post-eccentric exercise compared to all other groups (2.2-fold vs. SA/SED, 1.9-fold vs. SA/EX,

1.5-fold vs. LM-111/SED; all P<0.05; Figure 4.2A). A trend for an increase was observed in

LM-111/SED group compared to SA/SED (1.45-fold, P=0.076; Figure 4.2A). Expressing the data as total number of satellite cells per field of view, however, reveals a significant increase

(P<0.05) in LM-111/SED (17.7 cells/field) versus SA/SED (12.8 cells/field of view) (data not shown). Pax7 and Ki67 co-immunolabeling was used to quantify the number of proliferating satellite cells in young mice. The percentage of satellite cell expressing Ki67 was significantly higher in LM-111/EX (1.35%) as compared to SA/SED (0.5%), Saline/EX (0.8 %), and LM-

111/SED (0.57%; all P<0.05; Figure 4.2C). In young mice, we evaluated embryonic myosin heavy chain as a marker for newly synthesized myofiber during muscle regeneration (8). We noticed that the majority of eMHC+ myofibers were small in diameter, characteristic of newly formed myofibers 24 hours post-eccentric exercise (Figure 4.2F). Consistent with the satellite cell data, muscles injected with LM-111 demonstrated a significantly higher number of eMHC+

90 myofibers 24 hours after downhill running compared to all other groups (P<0.05) (Figure 4.2E). eMHC+ fibers were not detected in SA/SED.

LM-111 Directly Enhances Myoblast Proliferation

To determine the extent to which LM-111 could directly affect satellite cell proliferation, we treated myoblasts with LM-111 in culture. Flexcell plates consisting of silicone were coated with a thin layer of PDMS gel to mimic the microenvironment of young muscle in vivo (12 kPa).

The surface of the gels incorporated LM-111 in increasing concentrations or collagen as control.

Equal numbers of primary myoblasts were seeded on each well, and the total number of myoblasts was quantified 144 hours after plating. We observed 1.55 and 1.57-fold increases

(P<0.05) in myoblast quantity on gels coated with 10g/ml and 50g/ml of LM-111 compared to collagen coated plates, respectively, with no LM dose-effect (Figure 4.3A and B). A non- significant 1.35-fold change was observed with 30 µg/ml LM-111 treatment (P=0.097). From these observations, it was concluded that LM-111 directly increased myoblast proliferation, which may provide the basis for the elevation in baseline satellite cell content following LM-111 injection prior to injury.

LM-111 Does Not Increase mMSC Content, but Alters mMSC Gene Expression, in Young Mice

To better understand the mechanism by which LM-111 increased satellite cell quantity and muscle regeneration following eccentric exercise-induced damage, we studied the effects of

LM-111 injection on mMSCs. We focused on mMSCs given our previous data suggesting that mMSCs are sensitive to the extracellular niche and display different profiles of cell surface markers in the presence of gelatin or laminin in vitro (59). We first quantified the content of mMSCs in GAS complex of each mouse. No significant differences were found between groups for either the total number (Figure 4.4A), percentage of Sca-1+CD45- mMSCs (Figure 4.4B), or

91 number per mg of muscle (Figure 4.4C) after FACS. MSCs are known to respond to their microenvironment and promote tissue regeneration through the secretion of a variety of paracrine factors (38, 48); therefore, we analyzed the expression of genes involved in the mMSC secretome. The expression of interleukin-6 (IL-6) was significantly higher (2.64 fold) in SA/EX as compared to LM-111/Ex (P<0.05) (Figure 4.5A). A significant decrease in the expression of tumor necrosis factor-α (TNF-α), interleukin-1 receptor antagonist (IL-1rn) and interleukin-10

(IL-10) was observed with LM-111 injection (main effect, all P<0.05) (Figure 4.5B-D). The expression of hepatocyte growth factor (HGF), a potent activator of satellite cells (40), was significantly upregulated in mMSCs from LM-111/EX compared to SA/EX (1.5-fold, P<0.05)

(Figure 4.5E).

LM-111 Prevents Muscle Force Loss in Aged Skeletal Muscle

Body weight was significantly lower in LM-111-injected mice (main effect, P<0.05;

Figure 6B) and decreased with time in both groups (P<0.05; Figure 4.6B). No significant group differences in absolute or relative muscle weight were observed following eccentric exercise

(Data not shown).

Grip strength was used as a non-invasive measure of muscle function during the course of the post-injection period. Maximum grip strength was significantly higher in LM-111 injected mice (P<0.05; Figure 4.6D), while a trend for an increase in average grip strength was observed in LM-111 treated mice (p=0.052; Figure 4.6C). The changes observed with LM-111 injection were primarily due to an attenuation of the decrease in strength post-injection.

Satellite Cell Content in Old Mice Injected with LM-111

Satellite cells were quantified 24 hours following downhill running in both LM-111 and saline-injected mice. Satellite cell number was not altered with either LM-111 injection or

92 eccentric exercise in aged mice; however, a trend for an increase was observed in LM-111/EX

(P=0.09; Figure 4.7).

LM-111 Injection Alters the Function but not Quantity of mMSCs in Aged Skeletal Muscle

mMSC quantity was evaluated by FACS in both LM-111- and saline-injected muscles 24 hours following the acute downhill running protocol. Neither LM-111 nor downhill running increased the quantity of mMSC from aged skeletal muscle (Figure 4.8A and B). mMSCs isolated 24 hours post-downhill running were evaluated for the expression of genes implicated in inflammation, satellite cell activation and ECM remodeling. A trend for a decrease in IL-6 expression was observed in downhill run mice (main effect P=0.069; Figure 4.8C). In saline injected mice, TNF-α expression was significantly decreased with downhill running (P<0.05;

Figure 4.8D); while in downhill run mice, TNF-α expression trended higher in LM-111-injected mice (p=0.066; Figure 4.8D). Expression of FGF2 was significantly decreased in saline-injected mice exposed to downhill running (P<0.05; Figure 4.8E); while in LM-111-injected mice, fibroblast growth factor 2 (FGF2) expression increased post-downhill running (P<0.05; Figure

4.8E). In downhill run mice, FGF2 expression was significantly higher with LM-111 injection

(P<0.05; Figure 4.8E). No significant differences were observed in insulin growth factor (IGF-1) or (α1 type collagen) Collagen 1α expression (Figure 4.8F and G).

4.5 DISCUSSION

Previous studies have demonstrated the benefits of LM-111 therapy in mouse models of muscular dystrophy (21, 49-51, 60); however, the use of LM-111 therapy to enhance the muscle regenerative response to muscle damage in both young and old has not been previously examined. In the present study, we demonstrate that LM-111 injection increased the quantity and proliferation of satellite cells, enhanced the formation of new fibers and altered the expression of genes involved in the inflammatory and growth response in mMSCs in young mice. Furthermore, in old mice, LM-111 injection trended to increase the quantity of satellite cells and induce alterations in gene expression in mMSCs that favor myogenesis following eccentric exercise.

Together, these data support the hypothesis that LM-111 therapy can enhance the regenerative response to muscle damage in both young and old muscle.

Laminin is a primary component of the extracellular matrix (ECM) in skeletal muscle which surrounds the myofibers and forms a niche for muscle-derived stem cells. (18) In the present study, we found injected LM-111 to be present throughout the muscles of injected mice 7 days post-injection, and appeared to be localized to its natural location surrounding the myofibers. This is in agreement with previous studies that have detected LM-111 in injected muscles as long as 28 days following injection (49, 50, 60). In addition to diffuse LM-111 localization surrounding the myofibers, we also observed punctate areas of LM-111 that seemed to co-localize with Pax7+ satellite cells. Given that satellite cells highly express α7 integrin (20,

45), the primary LM-111 receptor (16), this observation suggests that LM-111 may have an intrinsic ability to migrate to areas of high receptor expression due to its affinity for α7 integrin

(44), and in this manner have direct effects on satellite cell function.

Previous studies have demonstrated that laminin is essential for normal myogenesis in vivo (30), and can affect myoblast adhesion, proliferation, and differentiation in vitro (22, 36). In the present study we demonstrated that in young mice, satellite cell quantity was increased only in LM-111 injected mice following eccentric exercise, and this response was due to increased satellite cell proliferation. These findings are in agreement with previous studies showing increased satellite cell quantity when LM-111 treatment was used in different mouse models of muscular dystrophy (21, 50, 60). To further investigate the direct effects of LM-111 on satellite cell regulation, we exposed primary myoblasts to different concentrations of LM-111 in vitro.

Treatment with 10 and 50 µg/ml of LM-111 resulted in a significant expansion of myoblast quantity as compared to collagen treatment. These data are in agreement with previous studies that have shown increase of myoblast proliferation with LM-111 addition in culture (21, 54). We did not observe a dose-response to LM-111 treatment suggesting that we had maximized the response with our lowest dose of LM-111. Indeed, a previous study by Silva-Barbosa and colleagues demonstrated that myoblasts stimulation with LM-111 treatment in vitro was maximized 10 µg/ml (54). The maximal increase in cell quantity of 20% in the study by Silva-

Barbosa and colleagues (54) is similar to the ~50% increase observed in the present study.

Furthermore, in the present study, we only quantified myoblast number at 6 days after plating. It is possible that a dose-response exists transiently at earlier stage of myoblast growth and disappears after the initial stages of myoblast activation.

Similar to the satellite cell response, we found an increased the number of eMHC+ myofibers after eccentric exercise in young LM-111-injected mice compared to saline-treated controls. Previous studies have reported eMHC+ myofibers as early as 4 days post injury (50, 52).

To the best of our knowledge, this is the first report of eMHC+ myofibers, a marker of new fiber

95 synthesis, 24 hours following muscle damage. The early detection of eMHC+ myofibers in the present study may be due to the mild and physiological amount of injury following eccentric exercise (downhill running versus cardiotoxin). Indeed, in response to cardiotoxin, the number of eMHC+ fibers is much higher at later time points post-injury (35, 52) which may be due to a prolonged inflammatory response. A second, intriguing explanation is the possibility that LM-

111 treatment directed a sub-population of activated myoblasts to rapidly differentiate. Data from previous in vitro studies have demonstrated that LM-111 facilitated myotube formation in a dose-dependent manner (58), and that myotube formation was decreased in the absence of LM-

111 treatment (55). Furthermore, LM-111 treatment in vivo increased the quantity of eMHC+ myofibers at both 4 and 10 days post injury compared to saline-injected mice (50, 60). Taken together, these data suggest that LM-111 can promote myogenesis directly through increasing the proliferation of satellite cells and by promoting myoblast differentiation.

Detrimental alterations in the aged muscle microenvironment, including the accumulation of collagen (1), are associated with impaired satellite cell function in response to injury (53).

Collagen accumulation and satellite cell dysregulation are also associated with muscular dystrophy (17, 42). Given the beneficial effects of LM-111 therapy on dystrophic mouse muscle, we hypothesized that LM-111 may improve the satellite cell response to muscle damage in aged mice. To allow for maximal incorporation of LM-111 into the muscle ECM and optimal preconditioning of the muscle microenvironment, we examined the acute response to downhill running 3.5 weeks post-injection. Numerous previous studies have demonstrated that LM-111 is still present in muscle at this time following injection (49, 50, 60). A trend for an increase in satellite cell quantity was observed in old mice treated with LM-111 24 hours following eccentric exercise, while no such changes were observed in saline treated mice. These data

96 suggest that LM-111 may partially restore the regenerative capacity of satellite cells in aged muscle.

It has been shown that ECM proteins play pivotal roles in myogenesis through the interaction with their receptors (58). The α7β1 integrin provides an important link between laminin (mostly laminin 1 and laminin 2) in the basement membrane and the inner actin cytoskeleton (4).. Increase of α7 integrin expression with either LM-111 injection or LM-111 transgenic overexpression in different mouse models of muscular dystrophy has been widely observed in previous studies (19, 21, 49, 60), yet the increase of α7 integrin expression is due to simply increased total amount of the protein or activating unbinding α7 integrin is still unclear.

In a recent study, Van Ry and colleagues observed increased sarcolemmal localization of α7 integrin after LM-111 treatment (60), suggesting that LM-111 injection may increase the binding sites for excessive unbinding α7 integrin located inside myofiber. It has been shown previously that activating excessive α7 integrin inside muscle fiber induced higher level of α7 integrin (33).

In the present study, we did not observe any increase in either α7A or α7B integrin expression

(Figure 4.9). We may have been unable to detect an increase in α7 integrin levels in the present study due to the different mouse models. Although α7β1 integrin is the predominant LM-111 receptor, we cannot exclude the possibility that injected LM-111 can also bind to the dystrophin- glycoprotein complex (DGC), the other receptor of LM-111 in skeletal muscle (9). In skeletal muscle of mdx mice, α7 integrin is the only receptor that LM-111 can bind due to the absence of dystrophin, whereas in healthy muscle tissue, LM-111 may bind to DGC after injection. As a result, injection of LM-111 into dystrophic muscle may result in an upregulation of α7 integrin because DGC is lacking which is not the case in young healthy or old muscle. Inconsistent with our previous finding (59),

To further examine the potential mechanisms that are responsible for improved muscle regeneration in LM-111 treated animals, we investigated both the quantity and gene expression of various cytokines and growth factors from mMSCs isolated from both LM-111 and saline- injected sedentary mice or 24 hours post-eccentric exercise. Given our previous data demonstrating that mMSCs accumulate in mice that transgenically overexpressed α7 integrin (35,

59), and the elevation of α7β1 integrin expression has been suggested as primary mechanism responsible for the benefits of LM-111 supplementation in mouse models of muscular dystrophy mice (49, 60), we hypothesized that LM-111 administration would increase mMSCs quantity in wildtype mice by increasing levels of α7 integrin. However, since we did not observe an increase in α7 integrin levels in response to LM-111 administration in both young and old mice, it is not surprising that mMSCs content did not increase in both young and aged animals. The different age and strain of mice used in this study may explain this discrepancy from our previous study

(46).

To gain a better understanding of mMSC function, we isolated mMSCs and evaluated the genetic expression of a variety of growth factors and cytokines. MSCs are clearly responsive to their microenvironment as previous studies have demonstrated differences in the secretion pattern of cytokines and growth factors when MSCs are cultured on different ECM substrates in vitro (47). Previously, we have observed alterations in the expression of cell surface markers from mMSCs cultured on laminin compared to gelatin (59), and have demonstrated that growth on laminin and exposure to mechanical strain promote secretion of factors that enhance vasculogenesis (25) and myogenesis (De Lisio et al., in preparation). In the present study, LM-

111 suppressed expression IL-6, after acute bout of downhill running in young mice. Although we didn’t measure serum levels of IL-6, previous studies have consistently found that systemic

IL-6 is significantly elevated 24 hours after a single bout of downhill running (7, 26). Thus, the suppression of IL-6 in mMSCs in LM-111 treated muscles, may suggest that muscle damage in response to downhill running was decreased in these animals. Meanwhile, increased HGF expression, a growth factor primarily responsible for stretch-induced satellite cell activation (56), was observed with LM-111 injection post-eccentric exercise in young mice. The concomitant accumulation of satellite cells suggests that HGF produced by mMSCs might be a key factor responsible for enhanced muscle regeneration with LM-111 supplementation. In addition, we observed a decrease in the expression of inflammatory cytokines in LM-111 treated mice regardless of exercise condition, suggesting LM-111 injection may significant influence mMSC function. The precise mechanism underlying the regulation of mMSCs function by LM-111 therapy is unknown. We speculate that since the majority of our mMSCs are pericytes (59), and laminins are found in the basement membranes of endothelium and encasing pericytes in vessel walls (61), then one explanation is that LM-111 injection may be able to alter the microenvironment of mMSCs in the perivascular niche. This hypothesis is supported by in vitro data from our lab where we consistently observe that mMSCs grown on laminin-coated versus collagen-coated plates alter their morphology, MSC marker expression, and the profile of trophic factors released (25, 59).

mMSCs gene expression was also altered by LM-111 supplementation in aged muscle.

IL-6, a potent activator of satellite cell proliferation following muscle damage (37), was decreased in mMSCs taken from both saline- and LM-111 injected mice exposed to eccentric exercise. The lack of upregulation of IL-6 may indicate that mMSCs from aged muscle are dysfunctional and cannot contribute to muscle repair, and this defect cannot be overcome by treatment with LM-111. Furthermore, in the present study, expression of FGF2 was significantly

99 increased in downhill run mice given LM-111 injections, while it was significantly decreased in saline injected mice post-eccentric exercise. FGF2 and TNF-α have been suggested to have direct effects on satellite cell activation and proliferation (28, 31). In old mice, it was recently shown that FGF2 levels are increased in the aged satellite cell niche which leads to excess satellite cell activation and eventual loss of the quiescent satellite cell pool (11). In the present study, the increase in FGF2 and trend for an increase in TNF-α gene expression occurred post- eccentric exercise, a time when satellite cell activation and proliferation are beneficial for muscle repair, only in mMSCs from LM-111 injected mice. These data suggest that LM-111 supplementation may be an effective therapy for increasing production of myogenic factors in aged mMSCs.

The ultimate goal of any therapeutic for aged muscle is to restore the loss of muscle mass and function. In the present study, we observed a decrease in muscle force production with saline injection that was attenuated by LM-111 injection. A previous study reported that LM-111 increased muscle force in mdx muscle with a single intramuscular injection (21). The authors suggested that the increase of muscle force may be due to the presence of new mechanical links between the extracellular matrix and sarcolemma by LM-111 and α7 integrin binding (21). The lack of increase in α7 integrin levels in the current study, suggest that in aged muscle a different mechanism may be responsible, possibly via interaction with DGC as described above.

Taken together, the present study provides the first evidence that LM-111 injection enhances skeletal muscle repair following a single bout of eccentric exercise. Our data indicate that LM-111 enhances early phase of regenerative response to eccentric exercise-induced injury in both young and aged muscle perhaps by altering growth factor production in mMSCs. Future studies are necessary to determine the exact factors secreted from mMSCs that are regulated by

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LM-111, and the precise mechanisms by which LM-111 alters mMSCs function in response to eccentric exercise. This information will guide us towards a better understanding of the development of therapeutic agents that can effectively restore regeneration capacity and preserve muscle mass and function with age and disease.

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4.6 FIGURES, TABLE AND FIGURE LEGENDS

Table 4.1. The primers used for qPCR in this study.

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Figure 4.1. LM-111 is detectable in young skeletal muscle after injection. (A) Experimental design and timeline for examining muscle regeneration following LM-111 administration in young mice. (B) The presence of LM-111 was detected by immunofluorescence in gastrocnemius/soleus complexes injected intramuscularly with LM-111, but not with saline. FITC (green) = Laminin α1, 20X, scale bar = 10 m. (C) Representative images of a muscle cross-section at 20x (scale bar = 20 m) and 40X (scale bar = 10 m) magnification stained for Laminin α1 in green, Pax7 in red, and DAPI in blue. Arrows indicate the co-localization of LM-111 and Pax7.

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Figure 4.2. LM-111 supplementation improves the regenerative response following eccentric exercise-induced damage in young skeletal muscle. (A) Quantification of Pax7+ satellite cells as a percentage of total myonuclei in LM-111 or saline injected muscle 24 hours after completing an acute bout of eccentric exercise (EX) or remaining sedentary (SED). (B) Representative image at 40x magnification of Pax7/α7 integrin co-stain with Pax7 in green, α7 integrin in red and DAPI in blue. Scale bar = 10 m (C) Quantification of proliferating satellite cells (Pax7+/Ki67+) as a percentage of total Pax7+ cells. (D) Representative image at 40x magnification of Pax7/i67 with Ki67 in green, Pax7 in red and DAPI in blue. Scale bar = 10 m (E) Quantification of embryonic myosin heavy chain positive (eMHC+) myofibers as the total number per section. (F) Representative image at magnification of 20x of eMHC stain with eMHC in green and DAPI in blue. Scale bar = 20 m * P < 0.05 compared with SA/SED; # P < 0.05 compared with SA/EX; & P < 0.05 compared with LM-111/SED. n = 6/group. Values are presented as mean ± SEM.

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Figure 4.3. LM-111 stimulates myoblast proliferation in vitro. (A) Quantification of myoblast proliferation on different concentrations of LM-111 and Collagen determined by cell counting with hemocytometer. Cell quantities are expressed relative to Collagen treated plates. (B) Representative images taken 120 hours after plating on different substrates. * P < 0.05 compared to Collagen. n = 6 wells/treatment. Values are presented as mean ± SEM.

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Figure 4.4. LM-111 supplementation does not increase mMSC content in young skeletal muscle. (A) Quantitation of total number of Sca-1+CD45- mMSCs collected after FACS. (B) Quantitation of the percentage of Sca-1+CD45- mMSCs in total mononuclear cells. (C) Quantification of mMSC percentage normalized to muscle weight. n = 6/group. Values are presented as mean ± SEM.

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Figure 4.5. LM-111 supplementation alters mMSC gene expression in young skeletal muscle. qPCR analysis of gene expression in mMSCs collected immediately after FACS for all groups. qPCR analysis of gene expression in mMSCs collected immediately after FACS for all groups in young mice. The graphs represent ΔΔCt values expressed relative to SA/SED. (A-D) Relative inflammatory cytokines expression: IL-6, TNF-α, IL-1rn and IL-10. (E) Relative HGF expression. # P < 0.05 compared with SA/EX; § main effect for LM-111 treatment. n = 5-6/group. Values are presented as mean ± SEM.

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Figure 4.6. LM-111 prevents muscle force loss in aged skeletal muscle. (A) Experimental design and timeline for examining muscle regeneration following LM-111 administration in aged mice. (B) Total body weight was recorded weekly during the 3.5 week after LM-111 injection. (C-D) Both fore- and hind-limb grip strength was assessed every week. Absolute average or maximal peak tension force (g) is presented. n = 6/group. Values are presented as mean ± SEM. * P<0.05, time effect; § main effect for LM-111 treatment.

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Figure 4.7. LM-111 therapy and satellite cell content following eccentric exercise in aged skeletal muscle. (A) Quantification of Pax7+ satellite cells as a percentage of total myonuclei in aged muscle. n=6/group.

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Figure 4.8. LM-111 supplementation on mMSC content and gene expression from aged skeletal muscle. (A) Quantitation of the percentage of Sca-1+CD45- mMSCs in total mononuclear cells. (B) Quantification of mMSC percentage normalized to muscle weight. (C-G) qPCR analysis of gene expression in mMSCs collected immediately after FACS for all groups in aged mice. The graphs represent ΔΔCt values. * P < 0.05 compared with SA/SED; # P < 0.05 compared with SA/EX. n = 5- 6/group. Values are presented as mean ± SEM.

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Figure 4.9. α7 integrin expression in young skeletal muscle. (A) Analysis of α7A expression (α7 CDB 345) and α7B expression (α7 CDB 347) in the gastrocnemius/soleus complex. Western blots of total 7 integrin probed with polyclonal antibody against total α7A or α7B integrin. Fold changes relative to SA/SED are presented. n = 6/group. Values are presented as mean ± SEM.

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42. Morgan JE, and Zammit PS. Direct effects of the pathogenic mutation on satellite cell function in muscular dystrophy. Exp Cell Res 316: 3100-3108, 2010. 43. Motohashi N, Uezumi A, Yada E, Fukada S, Fukushima K, Imaizumi K, Miyagoe- Suzuki Y, and Takeda S. Muscle CD31(-) CD45(-) side population cells promote muscle regeneration by stimulating proliferation and migration of myoblasts. Am J Pathol 173: 781-791, 2008. 44. Nishiuchi R, Takagi J, Hayashi M, Ido H, Yagi Y, Sanzen N, Tsuji T, Yamada M, and Sekiguchi K. Ligand-binding specificities of laminin-binding integrins: a comprehensive survey of laminin-integrin interactions using recombinant alpha3beta1, alpha6beta1, alpha7beta1 and alpha6beta4 integrins. Matrix Biol 25: 189-197, 2006. 45. Pasut A, Oleynik P, and Rudnicki MA. Isolation of muscle stem cells by fluorescence activated cell sorting cytometry. Methods in Molecular Biology 798: 53-64, 2012. 46. Peister A, Mellad JA, Larson BL, Hall BM, Gibson LF, and Prockop DJ. Adult stem cells from bone marrow (MSCs) isolated from different strains of inbred mice vary in surface epitopes, rates of proliferation, and differentiation potential. Blood 103: 1662-1668, 2004. 47. Prewitz MC, Seib FP, von Bonin M, Friedrichs J, Stissel A, Niehage C, Muller K, Anastassiadis K, Waskow C, Hoflack B, Bornhauser M, and Werner C. Tightly anchored tissue-mimetic matrices as instructive stem cell microenvironments. Nat Methods 10: 788-794, 2013. 48. Ranganath SH, Levy O, Inamdar MS, and Karp JM. Harnessing the mesenchymal stem cell secretome for the treatment of cardiovascular disease. Cell Stem Cell 10: 244-258, 2012. 49. Rooney JE, Gurpur PB, and Burkin DJ. Laminin-111 protein therapy prevents muscle disease in the mdx mouse model for Duchenne muscular dystrophy. Proc Natl Acad Sci 106: 7991-7996, 2009. 50. Rooney JE, Gurpur PB, Yablonka-Reuveni Z, and Burkin DJ. Laminin-111 restores regenerative capacity in a mouse model for alpha7 integrin congenital myopathy. Am J Pathol 174: 256-264, 2009. 51. Rooney JE, Knapp JR, Hodges BL, Wuebbles RD, and Burkin DJ. Laminin-111 protein therapy reduces muscle pathology and improves viability of a mouse model of merosin- deficient congenital muscular dystrophy. Am J Pathol 180: 1593-1602, 2012. 52. Serra C, Tangherlini F, Rudy S, Lee D, Toraldo G, Sandor NL, Zhang AQ, Jasuja R, and Bhasin S. Testosterone Improves the Regeneration of Old and Young Mouse Skeletal Muscle. Journals of Gerontology Series a-Biological Sciences and Medical Sciences 68: 17-26, 2013. 53. Shavlakadze T, McGeachie J, and Grounds MD. Delayed but excellent myogenic stem cell response of regenerating geriatric skeletal muscles in mice. Biogerontology 11: 363-376, 2010.

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54. Silva-Barbosa SD, Butler-Browne GS, de Mello W, Riederer I, Di Santo JP, Savino W, and Mouly V. Human myoblast engraftment is improved in laminin-enriched microenvironment. Transplantation 85: 566-575, 2008. 55. Smyth N, Vatansever HS, Murray P, Meyer M, Frie C, Paulsson M, and Edgar D. Absence of basement membranes after targeting the LAMC1 gene results in embryonic lethality due to failure of endoderm differentiation. J Cell Biol 144: 151-160, 1999. 56. Tatsumi R, Liu X, Pulido A, Morales M, Sakata T, Dial S, Hattori A, Ikeuchi Y, and Allen RE. Satellite cell activation in stretched skeletal muscle and the role of nitric oxide and hepatocyte growth factor. Am J Physiol Cell Physiol 290: C1487-1494, 2006. 57. Uezumi A, Fukada S, Yamamoto N, Takeda S, and Tsuchida K. Mesenchymal progenitors distinct from satellite cells contribute to ectopic fat cell formation in skeletal muscle. Nat Cell Biol 12: 143-152, 2010. 58. Vachon PH, Loechel F, Xu H, Wewer UM, and Engvall E. Merosin and laminin in myogenesis; specific requirement for merosin in myotube stability and survival. J Cell Biol 134: 1483-1497, 1996. 59. Valero MC, Huntsman HD, Liu J, Zou K, and Boppart MD. Eccentric exercise facilitates mesenchymal stem cell appearance in skeletal muscle. PLoS One 7: e29760, 2012. 60. Van Ry PM, Minogue P, Hodges BL, and Burkin DJ. Laminin-111 improves muscle repair in a mouse model of merosin-deficient congenital muscular dystrophy. Hum Mol Genet 2013. 61. Yousif LF, Di Russo J, and Sorokin L. Laminin isoforms in endothelial and perivascular basement membranes. Cell Adh Migr 7: 101-110, 2013. 62. Yu H, and Talts JF. Beta1 integrin and alpha-dystroglycan binding sites are localized to different laminin-G-domain-like (LG) modules within the laminin alpha5 chain G domain. Biochem J 371: 289-299, 2003. 63. Zou K, Meador BM, Johnson B, Huntsman HD, Mahmassani Z, Valero MC, Huey KA, and Boppart MD. The alpha(7)beta(1)-integrin increases muscle hypertrophy following multiple bouts of eccentric exercise. J Appl Physiol (1985) 111: 1134-1141, 2011.

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CHAPTER 5

CONCLUSIONS AND FUTURE DIRECTIONS

The overall aims of these studies were to investigate the role of mMSCs transplantation and LM-111 supplementation in muscle repair and adaptation following eccentric exercise. mMSC transplantation effectively improved skeletal muscle repair following multiple bouts of downhill running by increasing satellite cell number and myonuclear content. The enhancement of repair likely contributed to increased myofiber growth and muscle strength following long- term eccentric exercise training. In agreement with our previous findings, mMSC transplantation alone did not confer any beneficial changes in skeletal muscle, suggesting the exercised microenvironment may be necessary to provide important cues to stimulate mMSCs.

LM-111 supplementation similarly improved muscle repair following eccentric exercise.

Satellite cell proliferation and accumulation were significantly increased with LM-111 supplementation 24 hours after a single bout of downhill running. Regulation of mMSC gene expression by LM-111, specifically decreased expression of pro-inflammatory cytokines and increased expression of HGF, may contribute to the enhancement of repair observed in these studies.

In conclusion, the results of these studies demonstrate that both mMSC and LM-111 supplementation can successfully facilitate muscle repair following injury associated eccentric exercise. Therefore, we believe that our results provide sufficient evidence for developing mMSC and LM-111-based therapies to improve muscle repair with deficiencies associated with aging and neuromuscular disorders.

Further studies are necessary to investigate the following questions:

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1. What are the precise mechanisms by which mMSCs contribute to the adaptive response

to eccentric exercise? If mMSCs indirectly contribute to this process by secreting a

variety of paracrine factors, what factors are necessary?

2. In the current studies we supplemented mice with mMSCs isolated from α7Tg skeletal

muscle. To what extent can WT mMSCs induce similar beneficial changes following

eccentric exercise? These studies are currently underway in our laboratory.

3. LM-111 regulates mMSC function in skeletal muscle. Therefore, to what extent can

mMSC and LM-111 supplementation synergistically enhance muscle repair following

eccentric exercise?

4. What is the extent to which mMSC and/or LM-111 supplementation can restore

regenerative capacity and subsequently reduce loss of muscle mass and function with

aging and disease?

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APPENDIX A

COMPARISON OF GENE EXPRESSION IN SKELETAL MUSCLE-DERIVED STEM

CELLS FROM WILDTYPE AND α7 INTEGRIN TRANGENIC MICE FOLLOWING

SEDENTARY AND POST-EXERCISE CONDITIONS

INTRODUCTION

The accumulation of mMSCs in skeletal muscle is dependent on the presence of the 7 integrin subunit. This observation raises the question of whether mMSCs from α7Tg muscle are intrinsically different from WT. Our preliminary PCR data has demonstrated that several genes are uniquely expressed in mMSCs isolated from α7Tg muscle compared to WT, including toll- like receptor 4 (TLR4). TLR4 gene expression was higher in mMSCs isolated from WT muscle compared to α7Tg muscle that was protected from injury and inflammation following eccentric exercise. Given the fact that TLR4 plays an important role in regulating the secretion of inflammatory cytokines and growth factors synthesized by MSCs (1, 2), we hypothesized that mMSCs from α7Tg muscle may display a different secretome. Therefore, we compared the gene expression of mMSCs isolated from both WT and α7Tg mice following sedentary and post- exercise conditions.

METHODS

Animals. Protocols for animal use were approved by the Institutional Animal Care and Use

Committee (IACUC) of the University of Illinois at Urbana-Champaign. α7Tg mice

(SJL/C57BL6: MCK- α7BX2) were produced at the University of Illinois Transgenic Animal

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Facility as described . All experiments were conducted at approximately the same time of day.

Animals were fed standard laboratory chow and had access to water ad libitum.

Exercise protocol and Tissue Harvest. Five weeks old female WT and α7Tg mice remained at rest (basal conditions) or completed a single downhill running exercise (-20o, 17 m/min, 30 min).

Speed on the treadmill (Exer-6M, Columbus Instruments, Columbus, OH) was gradually increased from 10 to 17 m/min during a 2 min warm-up period. Mice were euthanized via carbon dioxide asphyxiation 24 hours post exercise. Sedentary WT and α7Tg basal controls were euthanized at the same age. Thus, the four groups compared were: WT/SED, WT/EX, α7Tg/SED and α7Tg/EX (n = 3/group). Twenty-four hours following the eccentric exercise, all mice were euthanized via carbon dioxide asphyxiation. The gastrocnemius-soleus complexes were rapidly dissected and used for fluorescence-activated cell sorting (FACS) to isolate Sca-1+CD45- cells.

Isolation of Sca-1+CD45- Cells from Skeletal Muscle. Sca-1+CD45- cells were isolated using flow sorting technique, as previously described (214). Briefly, gastrocnemius muscle from each animal was dissected along with soleus as a complex to retain Sca-1+CD45- cells found within the vascular niche primarily in the perimysium. After enzymatic digestion of the muscle tissue, filtered samples were incubated on ice with anti-mouse CD16/CD32 (1 μg/106 cells)

(eBioscience, San Diego, CA) for 10 min in order to block non-specific Fc-mediated interactions.

Following the block cells were stained in a cocktail of monoclonal anti-mouse antibodies Sca-1-

PE (600ng/106 cells) and CD45-APC (300 ng/106 cells) (eBioscience, San Diego, CA) diluted in

2% FBS in PBS. An aliquot of cells was used to determine total mononuclear cell content by manual counts with hemocytometer prior to FACS. Fluorescence activated cell sorting (FACS) was performed using an iCyt Reflection System (Sony Biotechnology Inc. Champaign, IL), located at Carle Hospital (Urbana, IL). Negative and single-stained controls were used to

120 establish gates. The resulting data were analyzed using WinList 3D (Verity Software House).

Sorted cell suspensions were centrifuged immediately after sorting and each cell pellet was lysed in 300 μl of RLT buffer (RNeasy Micro Kit). Cell lysates were then frozen for further gene expression analysis.

RNA extraction and RT-PCR. Total RNA was extracted from frozen sorted cells using the

RNeasy Micro Kit and QIAshredder spin columns according to the manufacturer's instructions

(Qiagen, Valencia, CA). The RNA concentrations and purities were analyzed spectrophotometrically by Synerg H1 Hybrid Multi-Mode Microplate Reader (Biotek, Winooski,

VT). The reverse transcriptase reaction was performed using first strand cDNA synthesis kit

Inc, Grand Island, NY) according to manufacturer’s instructions. Briefly, all stock primer/probe mixes were prediluted together in Tris–EDTA buffer to give 0.2x concentration. The preamplification reagent comprised 25 µL TaqMan PreAmp Master Mix and 12.5 µL pooled primers mix to 0.2x concentration. After adding 12.5 µL cDNA, the tubes were placed in a PCR thermal cycler and incubated at 95°C for 10 min. Samples were preamplified for 14 cycles. The preamplified samples were then diluted 1:20 with Tris–EDTA buffer. cDNA was stored at -20oC until qPCR analysis. Gene expression was individually quantified for all genes of interest using a qRT-PCR machine (7900HT Fast Real-Time PCR system and SDS Enterprise Database) and analyzed using the CT method. The genes we assessed are presented in Table A.1. Expression of GAPDH was used as the housekeeping gene.

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RESULTS

The Expression of Inflammatory Cytokines Was Suppressed in mMSCs Isolated From α7Tg

Mice Post Exercise

mMSCs isolated 24 hours post-downhill running from both WT and α7Tg mice were evaluated for the genetic expression of key members of the mMSC secretome that have been implicated in inflammation, satellite cell activation and proliferation. A significant decrease of

IL-6 expression was observed in mMSCs isolated from α7Tg/EX compared to α7Tg/SED group

(P<0.05; Figure A.1A). While TNF- expression was increased in WT/EX compared to

WT/SED and 7Tg/SED, a significant decrease in TNF-α expression was observed in α7Tg/EX

(P<0.05; Figure A.1B). While no significance was found in IL-12b gene expression, a trend for decrease was observed α7Tg/EX compared to WT/EX (P=0.079, Figure A.1C).

IL-13 expression was significantly elevated in mMSCs isolated from α7Tg/SED compared to WT/SED group (P<0.05, Figure A.1D). mMSCs isolated from both WT/SED and

α7Tg/SED had significantly lower IL-13 expression compared to α7Tg/SED, no matter of mice background (P<0.05, Figure A.1D). No difference exists between the mMSCs isolated from

WT/EX and α7Tg/EX.

The Expression of Growth Factors was Increased in mMSCs Isolated From α7Tg Mice Post

Exercise

The gene expression of HGF and EGF was higher in mMSCs isolated from α7Tg/24hr

PEx compared to all other groups (P<0.05, Figure A.2A and B).

CONCLUSION

The results from these experiments provide evidence that mMSCs isolated from α7Tg muscle following eccentric exercise uniquely express genes associated with inflammation,

122 satellite cell activation and proliferation. Pro-inflammatory cytokine gene expression is lower and growth factor gene expression is higher in mMSCs isolated from α7Tg muscle post-eccentric exercise. These changes may dramatically alter the potential for mMSCs to contribute to muscle repair and growth following exercise.

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FIGURES, TABLE AND FIGURE LEGENDS

Table A.1. The primers used for qPCR in this study.

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Figure A.1. The gene expression of inflammatory cytokines was suppressed in mMSCs isolated from α7Tg mice post exercise. qPCR analysis of gene expression in mMSCs collected immediately after FACS for all groups. The graphs represent ΔΔCt values expressed relative to WT/SED. (A-D) Relative inflammatory cytokines expression: IL-6, TNF-α, IL-12b and IL-13. (E) Relative HGF expression. * P < 0.05 compared with Saline/SED; # P < 0.05 compared with Saline/EX; & P < 0.05 compared with α7Tg/SED. n = 3/group. Values are presented as mean ± SEM.

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Figure A.2. The gene expression of growth factors was increased in mMSCs isolated from α7Tg mice post exercise. qPCR analysis of gene expression in mMSCs collected immediately after FACS for all groups. The graphs represent ΔΔCt values expressed relative to WT/SED. (A-B) Relative HGF and EGF expression. * P < 0.05 compared with WT/SED; # P < 0.05 compared with WT/24hr PEx; & P < 0.05 compared with α7Tg/SED. n = 3/group. Values are presented as mean ± SEM.

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REFERENCES

1. Wang Y, Abarbanell AM, Herrmann JL, Weil BR, Manukyan MC, Poynter JA, and Meldrum DR. TLR4 inhibits mesenchymal stem cell (MSC) STAT3 activation and thereby exerts deleterious effects on MSC-mediated cardioprotection. PloS one 5: e14206, 2010. 2. Waterman RS, Tomchuck SL, Henkle SL, and Betancourt AM. A new mesenchymal stem cell (MSC) paradigm: polarization into a pro-inflammatory MSC1 or an Immunosuppressive MSC2 phenotype. PloS one 5: e10088, 2010.

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APPENDIX B

Prior Work Regarding Effects of 7 Integrin on Eccentric Exercise-Induced Skeletal Muscle Hypertrophy

THE 71 INTEGRIN INCREASES MUSCLE HYPERTROPHY FOLLOWING MULTIPLE BOUTS OF ECCENTRIC EXERCISE*

Kai Zou1,2, Benjamin M. Meador1, Brian Johnson1, Heather D. Huntsman1,2, Ziad Mahmassani1,2,

M. Carmen Valero1,2, Kimberly A. Huey3and Marni D. Boppart1,2

Reprinted from Journal of Applied Physiology. Zou K, Meador BM, Johnson B, Huntsman HD, Mahmassani Z, Valero MC, Huey KA, Boppart MD. The α₇β₁-integrin increases muscle hypertrophy following multiple bouts of eccentric exercise, 2011, doi: 10.1152/japplphysiol. 00081.2011, with the permission of American Physiological Society.

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Abstract

Mechanical stimuli increase skeletal muscle growth in an mTOR- and p70S6K-dependent manner. It has been proposed that costameric proteins at Z bands may sense and transfer tension to these initiators of protein translation, but few candidates have been identified. The purpose of this study was to determine if a role exists for the α7 integrin in the activation of hypertrophic signaling and growth following eccentric exercise training. Five wk old, wild type (WT) and

α7BX2 integrin transgenic (α7Tg) mice were randomly assigned to one of two groups: 1) sedentary (SED) or 2) exercise training (EX). Exercise training consisted of downhill running 3 sessions/wk for 4 wk (-20°, 17 m/min, 30 min). Downhill running was used to induce physiological mechanical strain. Twenty-four hr following the final training session, maximal isometric hindlimb plantarflexor force was measured. Gastrocnemius-soleus complexes were collected for further analysis of signaling changes, including AKT, mTOR and p70S6K, and muscle growth. Although an increase in p70S6K activity was noted in WT/EX, no significant increases in CSA or force were observed in WT/EX compared to WT/SED. AKT, mTOR and p70S6K activation was higher, and whole muscle hypertrophy, relative muscle weight, myofibrillar protein, and force were significantly elevated in 7Tg/EX compared to 7Tg/SED.

A marked increase in average myofiber CSA was observed in 7Tg/EX compared to all groups.

Our findings demonstrate that the α7β1 integrin sensitizes skeletal muscle to mechanical strain and subsequent growth. Thus, the 71 integrin may represent a novel molecular therapy for the treatment of disuse muscle atrophy.

Keywords: integrin, exercise training, hypertrophy, p70S6K

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Background

A single bout of eccentric exercise initiates ultrastructural degradation and an inflammatory response in human and animal skeletal muscle (5, 9, 13). However, repeated bouts of eccentric exercise simultaneously create beneficial changes in skeletal muscle, including resistance to further injury (12) and accumulation of myofibrillar proteins leading to increased fiber cross sectional area (20, 27, 40, 41). Results from prior studies suggest that exercise- induced muscle growth is optimal following eccentric versus concentric or isometric exercise

(20, 40, 42, 43), and mechanical strain-induced increases in hypertrophic signaling and protein synthesis may be the underlying basis for this adaptation (22, 34, 49).

Mechanical stimuli can increase p70S6K phosphorylation in a rapamycin-sensitive and mTOR-dependent manner (4, 17, 25, 26, 30, 37), leading to increased initiation of protein translation in skeletal muscle. Elevated translational efficiency during exercise is not dependent on systemic factors or nutrients (insulin, IGF-1), as stretch-stimulated protein accumulation and growth can occur in cultured myotubes in the complete absence of systemic factors (48, 49) and load-induced increases in p70S6K phosphorylation and growth can occur in a phosphoinositide

3-kinase- and protein kinase B (PKB/AKT)-independent manner (22, 26, 37). In addition, levels of growth hormone and insulin-like growth factor (IGF-1) in the blood do not always correlate with enhanced protein synthesis in humans following eccentric exercise (51). It is clear that mTOR and p70S6K are intricately involved in the initiation of growth with mechanical strain

(21, 22), yet the upstream regulators of these signaling molecules are not well defined. Thus, it is important to identify intrinsic factors in muscle that have the potential to sense tension and convert mechanical signals into molecular events that regulate growth.

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West et al. (50) provides a concise overview of the limited candidate molecules that have the potential to sense and transmit tension across the sarcolemma, including phosphatidic acid

(PA) and/or costameric proteins such as focal adhesion kinase (FAK). PA, a lipid second messenger generated by the enzymatic activation of phospholipase D at the sarcolemma, is synthesized in tibialis anterior muscle in response to ex vivo eccentric contractions and addition of the PLD-PA inhibitor 1-butanol can sufficiently block downstream p70S6K activation (23,

39). FAK phosphorylation on tyrosine 397 is increased in response to mechanical loading upon transient transfection and this occurs prior to p70S6K activation and increased fiber growth (14,

28). These studies have shed light on the mechanisms responsible for p70S6K-mediated growth in response to mechanical strain. However, PA, FAK, and other unidentified costameric proteins may work together in a coordinated fashion to regulate p70S6K signaling and all of the players in this complex have yet to be defined.

Integrins are heterodimeric transmembrane receptors that allow cells to adhere to the extracellular matrix, providing the opportunity for cellular stabilization and the ability to transmit mechanical and chemical information from the outside to the inside of the cell (45). The α7β1 integrin is highly expressed in skeletal muscle and provides an important link between laminin in the basement membrane and the inner actin cytoskeleton at the costamere (3). Congenital myopathies caused by mutations in the α7 gene (ITGA7) in humans confirm a critical role for α7 integrin in skeletal muscle (19), and enhanced expression of the α7β1 integrin in skeletal muscle of mice with a severe form of muscular dystrophy (mdx/utr-/-) slows development of muscle pathology and markedly extends longevity (11). Although the precise mechanism by which the

7 integrin protects against muscle disease is not known, p70S6K phosphorylation is preferentially increased and fiber hypertrophy has been observed in mdx/utr-/- mice

131 overexpressing the 7 integrin transgene (8, 10). Thus, the 7 integrin may represent a novel intrinsic factor that can regulate growth in skeletal muscle.

We previously reported that MCK-driven expression of the 7 integrin in wild type mice suppresses hypertrophic signaling in the early time course (immediately, 3 hr) following an acute bout of eccentric exercise in the form of downhill running. However, our more recent study demonstrates the ability for the 7 integrin to accelerate myogenesis and fiber hypertrophy following a single bout of downhill running, with rapid accumulation of Pax7+ satellite cells and increased mTOR phosphorylation 1 day following exercise (32). The adaptations in 7Tg muscle were unexpected given the acute nature of the stimulus and suggest that the presence of the 7 integrin sensitizes muscle to mechanical strain-induced growth. In light of these results, we hypothesized that the presence of the 7 integrin would increase fiber hypertrophy and whole muscle size following repeated bouts of eccentric exercise. Therefore, the purpose of this study was to elucidate the extent to which the 7 integrin regulates hypertrophic signaling and growth in response to eccentric exercise training in the form of downhill running.

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Materials and Methods

Animals and Downhill Running Exercise

Transgenic mice overexpressing the 7BX2 integrin in skeletal muscle (SJ6/C57BL6:

MCK-α7BX2) were produced at the University of Illinois Transgenic Animal Facility as described (6, 11). Wild type (WT) littermates were used as controls. Mice were housed in a temperature-controlled specific pathogen-free animal room maintained on a 12:12 light-dark cycle at the animal facility at the University of Illinois. The animals were fed standard laboratory chow and water ad libitum. Five wk old female WT (n = 10) and α7Tg mice (n = 11) were randomly assigned to a sedentary (SED) or exercise training (EX) group. Exercise training consisted of downhill running 3 times/wk (Monday, Wednesday and Friday) for 4 wk (-20°, 17 m/min, 30 min) on a treadmill (Exer-6M, Columbus Instruments). Speed was gradually increased from 10 to 17 m/min during the first 2 min warm-up period for every running time. Protocols for animal use were approved by the Institutional Animal Care and Use Committee (IACUC) of the

University of Illinois at Urbana-Champaign and followed the animal care guidelines of the

American Physiological Society.

Contractile Force Measurements

Twenty-four hr following the final training session, maximal isometric hindlimb plantarflexor force was measured in the fasted state (fasting began 12 hr prior). Force was measured using a dual-mode lever system as described (35). Briefly, mice were fully anesthetized by i.p. administration of 100 mg/kg ketamine and 10 mg/xylazine prior to electrical stimulation of the sciatic nerve at 250 Hz for 1.5 sec. Muscle forces were collected using a servomotor and analog control unit (model 305C-LR, Aurora Scientific, Aurora, ON).

Stimulation was repeated nine times with a recovery period of 5 s, for a total of 10

133 measurements. The values of maximal force output were normalized to muscle mass (g/g).

Immediately following force measurements, gastrocnemius-soleus complexes from the stimulated limb were rapidly dissected and frozen in liquid nitrogen-cooled isopentane for immunohistochemistry studies, whereas muscles from the contralateral limb were directly frozen in liquid nitrogen for immunoblotting analysis.

Western Blot Analysis

Frozen gastrocnemius-soleus complexes from WT and α7Tg mice were manually ground with a mortar and pestle. Powdered tissue was homogenized in ice-cold buffer containing 20 mM

HEPES (pH = 7.4), 2 mM EGTA, 50 mM β-glycerophosphate, 1 mM dithiothreitol, 1 mM

Na3VO4, 1% Triton X-100, and 10% glycerol, supplemented with 10 µM leupeptin, 3 mM benzamidine, 5 µM pepstatin A, 1 mM phenylmethylsulfonyl fluoride and 10 µg/ml aprotinin.

The homogenates were rotated at 4oC for 1 h, centrifuged at 14,000 g for 15 min at 4oC, and supernatant was removed as the detergent-soluble fraction. Protein concentration was determined with the Bradford protein assay using BSA for the standard curve.

Equal amounts of protein (60 µg) were reduced, separated by SDS-PAGE using 6-8% acrylamide gels, and transferred to nitrocellulose membranes. Equal protein loading was verified by Ponceau S staining. Membranes were blocked in Tris-buffered saline (pH 7.8) containing 5%

BSA, incubated with the appropriate phospho-specific antibody (1:1000), and subsequently rinsed, blocked and reprobed with an antibody to determine total protein (Cell Signaling

Technology, Danvers, MA). HRP-conjugated anti-rabbit secondary antibodies (Jackson

ImmunoResearch Laboratories, Inc., West Grove, PA) were applied for 1 hr at 1:2000. The rat

α7B integrin cytoplasmic domain was detected by incubating membranes in purified α7 CDB

347 polyclonal rabbit anti-rat antibody (1:1000) for 1 hr (45). Bands were detected using

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SuperSignal West Dura Extended Duration Substrate (Thermo Scientific, Rockford, IL) and a

Bio-Rad ChemiDoc XRS system (Bio-Rad, Hercules, CA). Quantification was completed using

Quantity One software (Bio-Rad).

Assessment of Myofibrillar Protein

All procedures for the myofibrillar extractions were performed on ice or at 4°C as previously described (47). A small distal (30 – 60 mg) portion of sample was cut from the frozen muscle using a cold razor blade. The sample was quickly weighed on a balance. The weighed samples were manually ground with a mortar and pestle into powder and followed by homogenization in 20 volumes of a 250 mM sucrose buffer (250 mM sucrose, 100 mM KCl,

5mM EDTA, and 10 mM Tris, pH 6.8). Extracts were centrifuged at 3,000 g for 5 min, and the supernatant was discarded. The pellet was suspended in 10 volumes of a 0.5% Triton X-100 solution (150 mM KCl, 0.5% Triton X-100, and 10 mM Tris, pH 6.8), which is a modification of the solution described by Solaro et al. (38). The suspension was homogenized and centrifuged at

3,000 g for 10 min. The resultant pellet was rinsed with 10 volumes of wash buffer (150 mM

KCl and 20 mM Tris, pH 7.0) to remove excess Triton X-100 solution. The pellet was then suspended in 10 volumes of the wash buffer. Protein concentration was determined with the

Bradford protein assay using BSA for the standard curve.

Immunohistochemistry

Muscles were divided in half along the axial plane, embedded in OCT (Tissue-Tek;

Fisher Scientific), and transverse cryosections (8 µm) were cut from the center (3 sections per sample, separated by 100 µm) using a CM1850 cryostat (Leica, Wezlar, Germany). Sections were placed on frozen microscope slides (Superfrost; Fisher Scientific, Hanover Park, IL) and stored at -80oC prior to staining.

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To assess myofiber cross-sectional area (CSA), membranes were outlined by immunostaining for the α7 integrin. Slides were fixed in acetone for 5 min and blocked with PBS containing 5% bovine serum albumin (BSA) prior to incubation with 7CDB antibody (45).

FITC-labeled donkey anti-rabbit (1:100) (Jackson ImmunoResearch) was used to detect the integrin. The elimination of the primary antibody was included as a negative control in all immunohistochemistry and assessments were made only if background staining was absent.

Slides were mounted using Vectashield containing DAPI (Vector Laboratories) and examined blindly with a Leica DMRXA2 microscope (using the 20X objective). Images were captured using a Zeiss AxioCam digital camera and Axiovision software (Zeiss, Thornwood, NY, USA).

Fiber CSA was acquired using the advanced measurements component of Axiovision software on images obtained with a 20X objective. Mean CSA was obtained by measuring the areas of

1,000 fibers from each animal.

For evaluation of whole muscle hypertrophy, images were obtained from 7 integrin stained sections using a 5X objective. Due to the large size of the muscle cross sections from the center of the gastrocnemius, 10-20 images were obtained from each section and reconstructed into a single image with Adobe Photoshop, using transparency settings to overlay the images using morphological markers. The mean whole muscle CSA was then calculated for 3 sections/animal using Axiovision software.

Statistical Analysis

All averaged data are presented as the means ± SE. To determine significance, comparisons between groups were performed by two-way ANOVA to determine if an interaction effect was observed for group and time. Tukey (7 integrin protein, fiber CSA, whole muscle

CSA, body weight changes, muscle mass, total protein, myofibrillar protein) or the least

136 significant difference (LSD) (isometric force, signaling) analyses were performed when appropriate interaction or main effects were found. All calculations were performed with SPSS statistical software (16.0, Chicago, IL). Differences were considered significant at P < 0.05.

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Results

α7 Integrin is Not Altered in WT Mice Following Exercise Training. Total 7 integrin protein was increased 6-7-fold in 7Tg mice compared to WT, with no further enhancement observed in either group following the 4 wk training period (Fig. B.1A). Immunofluorescence analysis of muscle cross sections verified localization of endogenous and transgenic 7 integrin at the sarcolemma and sarcoplasm of 7Tg muscle fibers, with highly concentrated protein observed at the membrane in enlarged fibers (Fig. B.1B).

α7 Integrin Positively Influences Hypertrophic Signaling Following Eccentric Exercise Training.

AKT-mTOR-p70S6K signaling has been shown to play an integral role in the initiation of protein synthesis following mechanical stimuli (21, 22). Despite the repeated exposure to eccentric exercise, total amounts of AKT, mTOR and p70S6K protein were unaltered as assessed by both immunoblotting and ponceau S staining in both WT and 7Tg mice 24 hr following the

4 wk training period (Fig. B.2A & 2B). Although AKT and mTOR phosphorylation remained unchanged, a 50% (P<0.05) increase in p70S6K phosphorylation was observed in WT/EX compared to WT/SED (Fig. B.2B). Phosphorylation and activation of all three proteins were increased following training in 7Tg/EX compared to 7Tg/SED (AKT, 1.7-fold; mTOR, 2.0- fold; and p70S6K, 1.3-fold) (P<0.05) (Fig. B.2B).

α7 Integrin Promotes Myofiber and Whole Muscle Enlargement Following Eccentric Exercise

Training. Extensive analysis of muscle CSA in 1000 individual fibers per animal revealed marked area increases in 7Tg fibers compared to WT in both the sedentary and post-training conditions. In sedentary and trained WT muscle, fibers were predominantly small, with the majority measuring 0-500 or 500-1000 µm2 (Fig. B.3A). The presence of the 7 integrin in the muscle increases the proportion of large caliber fibers, with relatively few small fibers observed

138 in the sedentary state compared to WT/SED (Fig. B.3B). We specifically noted a robust elevation in the percentage of fibers ranging 2000 to 3000 µm2 in 7Tg muscle, both in the sedentary state and post-exercise (Fig. B.3C). An interaction effect was observed between group and time for the percentage of fibers ranging 2000 to 3000 µm2. A 12-fold increase was observed in 7Tg/EX compared to WT/EX and a 2.5-fold increase compared to α7Tg/SED (P<0.001).

Mean myofiber CSA was increased 2.1-fold in 7Tg/SED compared to WT/SED and was increased 40% in α7Tg/EX compared to α7Tg/SED (1790.3 ± 141.9 vs. 1280.8 ± 62.25) (P <

0.01) (Fig. B.3D). Whole muscle CSA was increased approximately 30% in 7Tg/EX compared to both WT/SED and WT/EX (P<0.05) (Fig. B.4B)

α7 Integrin Enhances Muscle Force Production Following Eccentric Exercise Training. No group differences in body weight or absolute muscle weight occurred during 4 wk of eccentric exercise training (Fig. B.5A, absolute muscle weight data not shown). Under sedentary conditions, muscle mass relative to body mass was similar between WT (5.69 ± 0.16) and α7Tg mice (5.48 ± 0.05). Relative muscle mass was higher in α7Tg mice compared to their sedentary counterparts after 4 wk of eccentric exercise training (P < 0.05) (Fig. B.5B), whereas, no significant change was found in WT mice after training. Myofibrillar protein content expressed relative to muscle weight (mg/mg) was increased 98% in 7Tg/EX compared to WT/EX and

2.3-fold compared to 7Tg/SED (P< 0.05) (Fig. B.5C). Consistent with the increase in myofibrillar content, a 25% improvement in maximal isometric force was apparent in 7Tg/EX compared to 7Tg/SED when normalized to muscle mass (g/g) (169.2 ± 12.1 vs. 210.8 ± 9.8) (P

< 0.05) (Fig. B.5D). Maximal isometric force was not different in WT/EX compared to WT/SED

(P = 0.55).

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Discussion

We recently reported that the α7 integrin accelerates myogenesis and myofiber hypertrophy following an acute bout of eccentric exercise (32), suggesting an intrinsic role for this structural protein in exercise-induced muscle growth. This study is the first to investigate the extent to which overexpression of the 7BX2 transgene increases hypertrophic signaling, fiber size and whole muscle growth following multiple bouts of downhill running. Whereas minimal changes were noted in WT muscle in response to downhill running exercise, myofiber CSA, whole muscle CSA, relative muscle weight, myofibrillar protein content, and maximal isometric force were significantly elevated in 7Tg muscle following training compared to sedentary counterparts. AKT, mTOR, and p70S6K were also activated in 7Tg muscle post-exercise and may underlie the beneficial adaptations observed in this study..

Downhill running can initiate a regenerative response in WT mouse soleus muscle (1), consistent with our recently acquired data (32). Resistance training can increase p70S6K signaling (34, 51) and endurance training can enhance mTOR phosphorylation (2) in human skeletal muscle. However, we are not aware of any studies that have examined hypertrophic signaling, myofiber size, muscle mass or force in mice or humans following downhill run training. The results provided here do not support the presence of hypertrophy in WT muscle with 4 wk of downhill run training. Significant increases in p70S6K phosphorylation and a shift toward larger caliber fibers indicate potential for adaptation to downhill running. Most studies in the literature do not show evidence of increased muscle strength until 6-12 wk eccentric resistance exercise training (18, 38). Thus, additional weeks of downhill run training might elicit a more optimal growth response in WT mouse muscle. On the other hand, adaptations to regular

140 running may preclude substantial increases in muscle growth and strength that would normally occur as a result of lengthening contractions.

In contrast to the lack of hypertrophy observed in WT muscle, parameters of signaling and growth were consistently increased in 7Tg muscle in response to multiple bouts of eccentric exercise. The mean myofiber CSA was approximately the same after repeated bouts (1790 um2) compared to a single bout of downhill running exercise (1734 um2) reported in our previous study (32), however, the distribution of fiber sizes was altered following training with a lower percentage of 1000-2000 um2 fibers and an increase in the percentage of fibers measuring 2000-

3000 um2. Whole muscle CSA was enhanced following training (16,176,580 um2) vs. an acute bout (14,013,766 um2). In addition, myofibrillar protein was substantially increased following training. Whereas 40-200% increases in plantaris weight are standard within a couple of weeks following synergistic ablation (36, 37), we do not observe similar dramatic increases in gastrocnemius-soleus weight (~11% increase). We would have predicted higher muscle weight and force in 7Tg muscle compared to WT muscle based on the marked increases in fiber CSA, whole muscle CSA and myofibrillar protein post-exercise. In addition, while p70S6K phosphorylation and mean myofiber CSA were elevated in 7Tg/SED compared to WT/SED, muscle weight and isometric force were not different between these groups. We cannot account for the discrepancies in 7Tg/SED. However, the relatively lower weight reported in this study for 7Tg/EX compared to other studies that use mechanical overload may result from alterations in muscle metabolism and tissue composition caused by endurance exercise. Loaded wheel running provides similar small changes in muscle weight (~20%) (29). Application of an appropriate weight bearing model such as synergistic ablation to 7Tg mouse muscle would likely resolve these discrepancies. Despite this limitation, it is clear that the 7 integrin can

141 significantly increase signaling, muscle fiber size and myofibrillar protein synthesis in response to downhill running. Future molecular therapies focused on the use of the 7 integrin to combat muscle atrophy will likely need to incorporate mechanical strain or alternative methods to activate the integrin and optimally restore function.

Skeletal muscle damage can occur as a result of eccentric exercise in both humans and animals (5, 9, 13), as manifested by immediate and delayed injury to the sarcolemma, disruption of the excitation-contraction (E-C) coupling complex, muscle soreness, and prolonged strength loss (13, 15). Whereas injury is clearly evident in WT muscle following a single bout of downhill running, muscle overexpressing the 7 integrin transgene is protected from membrane damage, deficits in isometric force, and macrophage accumulation (6, 32). Reduced injury in 7Tg muscle likely creates a healthy microenvironment that is conducive to growth, making it difficult to distinguish whether the 71 integrin primarily acts as a mechanosensor, directly converting tension into molecular events that initiate increased protein synthesis, or simply serves a permissive role in fiber growth. It is likely that 7 integrin tethers laminin to the actin cytoskeleton as an internal mechanism to resist mechanical force and injury and that increased

7 integrin activation in the form of increased adhesion has direct consequences for signaling events inside muscle. Interestingly, actin disruption with cytochalasin D has been shown to inhibit strain-induced increases in p70S6K phosphorylation in myotubes (21). Support for this hypothesis is also evident in a study by Klossner et al. (28) showing increased p70S6K activation following in vivo plasmid transfer of the integrin-associated focal adhesion kinase (FAK).

Whether the 7 integrin influences phosphatidic acid accumulation in skeletal muscle is not known, but worthy of further investigation (23, 39). Our upcoming in vitro experiments utilizing

7 integrin-overexpressing primary myotubes should be able to address the extent to which the

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71 integrin can become activated and influence FAK activation, PA synthesis, and/or the downstream p70S6K signaling response to mechanical strain.

Although we previously reported increases in 7 integrin RNA and protein following a single bout of eccentric exercise (7), we did not observe elevations in 7 integrin protein with multiple bouts in WT muscle. These data suggest that upregulation of the integrin following exercise may be transient and newly synthesized protein may be less important to the prevention of injury and sustained intracellular signaling than increased adhesion. However, we suspect that

7 integrin protein may have been upregulated in muscle post-exercise training, but not detected in this study as a result of the detergent used for extraction (Triton-X). We are currently examining 7 integrin protein in tissues extracted with SDS detergent to determine if 7 integrin is recovered in the cytoskeletal fraction as a result of increased adhesion. Increased amount and affinity of the 7 integrin to extracellular matrix and actin may serve to decrease protein degradation and contribute to net increases in protein synthesis following long-term resistance training (33, 50).

This study provides the first demonstration that the 71 integrin increases AKT-mTOR- p70S6K signaling and muscle hypertrophy following eccentric exercise training in mice. The

71 integrin may represent a critical intrinsic factor in muscle that initiates signaling events leading to efficient exercise-induced muscle growth. Further studies are necessary to elucidate the molecular events linking 7 integrin activation with protein synthesis in skeletal muscle following eccentric exercise.

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We would like to thank Dr. Charlotte Peterson, University of Kentucky, for her insightful comments regarding our work and providing the protocol to evaluate myofibrillar protein. This work was supported by grants from the Illinois Regenerative Medicine Institute, Ellison Medical

Foundation, Mary Jane Neer Foundation (UIUC), and Arnold O. Beckman Award (UIUC) (to

MDB).

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Figures and figure legends

Figure. B.1. Transgenic expression of the 7 integrin. A: Analysis of 7 expression (7 CDB 347) in the gastrocnemius-soleus complex. Representative immunoblots of total 7 integrin probed with polyclonal antibody against total 7B integrin. Fold changes relative to WT/SED are presented (n = 4- 5/WT; n = 6/7Tg). B: Representative immunostaining of 7 integrin in the sedentary state (SED) and 24 h following the final eccentric exercise training session (EX). *P < 0.05 compared with W/SED; # P < 0.05 compared with WT/EX. Data are presented as mean ± SE.

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Figure B.2. Eccentric exercise training increases activation of hypertrophic signaling in 7Tg muscle. Gastrocnemius-soleus complexes were collected 24 hr following the last bout of exercise training in the fasted state. A: Representative immunoblots of AKT, mTOR, and p70S6K phosphorylation and expression from WT and 7Tg muscle in the sedentary state (SED) and following exercise training (EX). B: (Top) Analysis of AKT phosphorylation (Ser473), (Middle) mTOR phosphorylation (Ser2448), and (Bottom) p70S6K phosphorylation (Thr389). Fold changes relative to WT/SED are presented (n = 4- 5/WT; n = 3-4/7Tg). *P < 0.05 compared with W/SED; # P < 0.05 compared with WT/EX; §P < 0.05 compared with 7Tg/SED. Data are presented as mean ± SE.

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Figure. B.3. Mean individual fiber cross sectional area is increased in 7Tg mice following 4 wk of eccentric exercise training. Cross sectional area (CSA) was measured in 1000 fibers from WT and 7Tg gastrocnemius-soleus complexes following exercise training. A: Distribution of fiber sizes in WT mice following 4 wk eccentric exercise training. B: Distribution of fiber sizes in 7Tg mice following 4 wk eccentric exercise training. C: Percentage of fibers with CSA between 2,000 to 3,000 m2 (n = 3-4/grp). D: Mean fiber CSA (n = 3-4/grp). * P < 0.05 compared with WT/SED; # P < 0.05 compared with WT/EX; §P < 0.05 compared with 7Tg/SED. Data are presented as mean ± SE.

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Figure. B.4. Whole muscle cross sectional area is increased in 7Tg mice following 4 wk of eccentric exercise training. Gastrocnemius-soleus cross sectional area (CSA) was measured in WT and 7Tg mice following exercise training. A: 10-20 images were obtained using a 5X objective and reconstructed into a single image. Representative image for 7Tg/EX at 4 wk post-training is shown. B: Mean whole muscle CSA (n = 3-4/grp). * P < 0.05 compared with WT/SED; # P < 0.05 compared with WT/EX. Data are presented as mean ± SE.

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Figure B.5. Muscle weight, myofibrillar protein and maximal isometric force are increased in 7Tg mice following 4 wk of eccentric exercise training. A: Changes in body weight during the training period. B: Wet gastrocnemius-soleus complex weight expressed relative to body weight (n = 4-6/WT; n = 4-5/7Tg). C: Changes in myofibrillar protein content expressed relative to muscle weight (n = 4-5/WT; n = 3-4/7Tg). D: Changes in average maximal isometric plantarflexor force (normalized to wet gastrocnemius-soleus complex weight) 24 hr following the last bout of eccentric exercise training in WT and 7Tg mice (n = 4-6/WT; n = 4-5/7Tg).§P < 0.05 compared with 7Tg/SED. Data are presented as mean ± SE.

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38. Norrbrand L, Pozzo M, and Tesch PA. Flywheel resistance training calls for greater eccentric muscle activation than weight training. Eur J Appl Physiol 110: 997-1005. 39. O'Neil TK, Duffy LR, Frey JW, and Hornberger TA. The role of phosphoinositide 3- kinase and phosphatidic acid in the regulation of mammalian target of rapamycin following eccentric contractions. J Physiol 587.14: 3691-3701, 2009 40. Roig M, O'Brien K, Kirk G, Murray R, McKinnon P, Shadgan B, and Reid WD. The effects of eccentric versus concentric resistance training on muscle strength and mass in healthy adults: a systematic review with meta-analysis. Br J Sports Med 43: 556-568, 2009. 41. Roig M, Shadgan B, and Reid WD. Eccentric exercise in patients with chronic health conditions: a systematic review. Physiother Can 60: 146-160, 2008. 42. Seger JY, Arvidsson B, and Thorstensson A. Specific effects of eccentric and concentric training on muscle strength and morphology in humans. Eur J Appl Physiol Occup Physiol 79: 49-57, 1998. 43. Seger JY and Thorstensson A. Effects of eccentric versus concentric training on thigh muscle strength and EMG. Int J Sports Med 26: 45-52, 2005. 44. Solaro RJ, Pang DC, and Briggs FN. The purification of cardiac myofibrils with Triton X-100. Biochim Biophys Acta 245(1) :259-262, 1971. 45. Song WK, Wang W, Sato H, Bielser DA, and Kaufman SJ. Expression of alpha 7 integrin cytoplasmic domains during skeletal muscle development: alternate forms, conformational change, and homologies with serine/threonine kinases and tyrosine phosphatases. J Cell Sci 106 ( Pt 4): 1139-1152, 1993. 46. Spangenburg EE. Changes in muscle mass with mechanical load: possible cellular mechanisms. Appl Physiol Nutr Metab 34: 328-335, 2009. 47. Thomason DB, Herrick RE, and Baldwin KM. Activity influences on soleus muscle myosin during rodent hindlimb suspension. J Appl Physiol 63(1):138-44 :138-144, 1987 48. Vandenburgh H and Kaufman S. In vitro model for stretch-induced hypertrophy of skeletal muscle. Science 203: 265-268, 1979. 49. Vandenburgh HH. Motion into mass: how does tension stimulate muscle growth? Med Sci Sports Exerc 19: S142-149, 1987. 50. West DW, Burd NA, Staples AW, and Phillips SM. Human exercise-mediated skeletal muscle hypertrophy is an intrinsic process. Int J Biochem Cell Biol 42: 1371-1375. 51. West DW, Kujbida GW, Moore DR, Atherton P, Burd NA, Padzik JP, De Lisio M, Tang JE, Parise G, Rennie MJ, Baker SK, and Phillips SM. Resistance exercise-induced increases in putative anabolic hormones do not enhance muscle protein synthesis or intracellular signalling in young men. J Physiol 587: 5239-5247, 2009.

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APPENDIX C

PROTOCOLS

Sca-1+CD45- mMSCs Isolation Protocol ...... 156 Laminin-111 Injection Protocol ...... 159 Immunohistochemistry Protocol (Dystrophin) ...... 160 Immunohistochemistry Protocol (F4/80) ...... 161 Immunohistochemistry Protocol (Laminin-1) ...... 162 Immunohistochemistry Protocol (Pax7/Dystrophin) ...... 163 Immunohistochemistry Protocol (Pax7/Ki67) ...... 165 Myofiber Cross Sectional Area Measurement Protocol ...... 167 Myonuclear Content Measurement Protocol ...... 168 Muscle Homogenization Protocol...... 169 Western Blotting Protocol...... 170

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Sca-1+CD45- mMSCs Isolation Protocol

Dissection:

1. Exercise mice on a treadmill set at 20 degree decline with a speed of 17m/min, for 30 min, 24 hours prior to excising the muscle 2. Sacrifice 2 mice 5 weeks old by CO2 inhalation 3. Using autoclaved dissection tools, dip the whole mouse with 70% ethanol and excise both gastroc/soleus complexes from each mouse under a hood 4. Rinse the muscles with HBSS + P/S (from one of the tubes that have 10ml HBSS + P/S) in a petri dish to wash off the hair and put in a 50ml falcon tube (in 1736 cabinet)which has HBSS + P/S to keep the muscles moist and free from bacteria 5. Keep the muscle tissue on ice 6. Try to maintain as sterile of an environment as you can 7. Transfer the muscles to a “clean stem cell” ice container before bringing them into the cell culture room

Mince Muscle and Dissociate Cells:

1. Begin warming Enzyme Solution 1 in the sterile water that you have placed in the 37 degree Incubator before starting dissection 2. Before starting your work under hood, wipe down everything inside the hood and anything that you will bring under the hood, with 70% ethanol 3. Rinse the muscles in a sterile 60mm petri dish with HBSS + P/S before mincing 4. Place two muscles at a time in a sterile 60mm petri dish and add a few drops of HBSS + P/S to keep the tissue moist (approximately 350ul per dish) 5. Using autoclaved, curved scissors, mince the muscle into a slurry 6. Add the DNAse and Collagenase to Enzyme Solution 1 that you have just taken out of the incubator. When taking out the DNAse, DO NOT up and down pipette. 7. Transfer all the minced tissue with an autoclaved spatula to a sterile 15ml falcon tube (Orange cap) and add 2.5ml (per mouse) of fresh pre-warmed Enzyme Solution 1 8. Use a micro pipette to rinse the dish, scissors, and spatula with the Enzyme Solution 1, make sure you get all of the cells into the tube 9. Incubate in a beaker with pre-warm water in 37 degree incubator for 30-45 min, or until the mixture is fine slurry and you cannot see large pieces of muscle. (every 10-15 min gently mix the solution by pulling it up and down in 10ml pipette under the hood) 10. The supernatant now contains the cells we are interested in. The remaining muscle and collagen fragments will settle to the bottom of the tube

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11. Transfer the solution with slurry to 50ml falcon tube (stem cell specific) and add a volume equal to the amount of Enzyme solution 1 of inhibition-Medium to the solution to facilitate filtration (sometimes you need to add more, add up to 30 for 3 mice) 12. Using two 50ml BD (stem cell specific) tubes labeled #1 and #2, strain the inhibited solution 2x’s with two 70um strainer and 2x’s with two 40um strainer. After the first filtration and approximately 15-20 ml of inhibition medium to further aid in filtration. Also, use 10ml pipette to push the solution through the filter slowly. Mix couple of times before each filtration. Use foam rack to hold the 50ml tubes. Also do all the experiment very inside the hood to avoid contamination! 13. Centrifuge the cells for 5 min at 450 RCF. After centrifuge, move the tube very carefully, because cells are very easy to re-suspend in buffer! At mean time, take out Fc receptor blocking solution and antibodies from refrigerator 14. Remove the supernatant under the hood with a pasteur pipette, DO NOT touch the cell layer, you will lose some cells 15. Re-suspend the cells in 1ml of 2% FBS in PBS, add 1ml (add drop by drop at the beginning to avoid the temperature shock), up and down sufficiently and finger vortex the cells to ensure they are evenly suspended in the solution 16. Transfer cells to 2ml eppendorf tube. Remove 20ul of cell suspension and place in a 0.5ml eppendorf tube on ice to use later for counting 17. Add 5ul of Fc receptor blocking solution on 1ml of cell suspension to achieve 0.5ug/ml 18. Incubate for 10min with blocking solution on ice

Antibody Staining:

1. After blocking, mix the cells again, take 50ul of cell suspension and put in a sterile 2.0ml eppendorf tube. This will serve as the unstained control. 2. After blocking, Centrifuge for 10min at 450 RCF (cold centrifuge in the main room in 3636) at 4 degree 3. Remove supernatant with blue tip. Re-suspend in appropriate volume. Then add the antibody mix to the solution to make up to certain volume in order to aliquot 100ul per eppendorf tube (2ml tubes with curved bottoms and sterile) (e.g. 8 aliquots, you need to add 764ul 2% FBS + 24ul PE + 12ul APC) 4. Finger vortex each tube to evenly distribute the cells in suspension 5. Incubate in the dark on ice for 1hour, mixing the samples every 15min 6. Combine all of the eppendorf tubes into 15ml falcon tube and add 3-4ml of 2% FBS in PBS. Centrifuge. 7. Re-suspend the pellet in 7-8ml of 2% FBS in PBS and put in appropriate tube for sorting. For each time, add 1ml FBS to the cell (drop by drop at the beginning). Add the rest amount with 5ml pipette. Repeat 2 times. 8. Add 300ul of 2% FBS in PBS to your control cells and put in the appropriate sorting tube. Label your tubes well

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9. Remove most of the supernatant with Pasteur pipette and take out the rest small amount with yellow tip. Add 2ml FBS in PBS in the cells, mix well (up and down many times) and transfer to the other sorting tube. 10. Remove 50ul of cell suspension and take this over to sorting facility as well. Later when we get back, we will spin down at 450 RCF for 5min, then plate in 3ml of media on a 35mm petri dish. Place in the left incubator. This will serve as your contamination control 11. Prepare 2-15ml falcon tubes with approximately 5ml of media for sorting. Keep everything on ice. Also take an additional 15ml tube with some 2% FBS in PBS over to the facility, just in case our cells are too concentrated.

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Laminin 111 Injection Protocol

1. Laminin 111 aliquots (36µl for each) are kept at -20 degree freezer. and let them thaw slowly in 4 degree overnight 2. Pour some TBS (50 mM Tris-HCL (pH 7.4), 0.15 M NaCl in PBS) through 40µm strainer to sterilize it 3. Before injection, animals are anesthetized with 2% isofluorane, administered by inhalation (about 30s) 4. Wipe the gastrocnemius muscle with 70% Ethanol. 5. Total of 100 µl of Laminin 111 solution or TBS is injected into each muscle. Two separate injections, each of 50 µL, were injected into the lateral and medial lobe of the gastrocnemius to ensure uniform distribution throughout the entire muscle.

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Immunohistochemistry Protocol– Centrally Located Nuclei, Cross Sectional Area and Myonuclear Content (Dystrophin)

Procedure Directions

Section Preparation 8 µm sections placed on the slide; Allow frozen slides to thaw and dry

Fix Place all slides in acetone - 10 minutes (in freezer) Remove from acetone, allow time to dry, then circle sections with immunopen

Wash & Rehydrate 1x PBS - 5 min, 3 times

Block 5% BSA - 20 min

Fabs in 5% BSA - 30 min 1:20

1st Primary Antibody Dystrophin (Mandra; Sigma) 1:100 dilution in 1% BSA - 30 min @ 37°C

Wash 1% BSA (in 1x PBS) - 5 min, 3 times

1st Secondary FITC anti-mouse 1:250 dilution in 1% BSA - 60 min Antibody place slides in drawer *In Dark*

Wash 1% BSA/PBS—5min

DAPI-5min

1% BSA/PBS—5min

Coverslip 1 drop of Vectashield over each section Apply coverslip Seal with nail polish Store in refrigerator for up to 1 year

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Immunohistochemistry Protocol – Macrophage Content (F4/80)

Procedure Directions

Section Preparation 8 µm sections placed on the slide; Allow frozen slides to thaw and dry

Fix Place all slides in acetone - 10 minutes (in freezer) Remove from acetone, allow time to dry, then circle sections with immunopen

Wash & Rehydrate 1x PBS - 5 min, 3 times

Block 10% Goat Serum (in 1x PBS) - 20 min

Fabs in 10% GS - 30 min 1:20

Primary Antibody F4/80 1:100 dilution in 10% GS - 60 min *Don’t Put On Negative Control - Leave wash on from last step*

Wash 1% GS (in 1x PBS) - 5 min, 3 times This solution is used for multiple steps and multiple solutions.

Secondary Antibody *In Dark* FITC anti-Rat 1:100 dilution in 1% GS - 60 min place slides in drawer

Wash 1% GS (in 1x PBS) - 5 min DAPI 1:20000 – 5 min 1% GS (in 1x PBS) - 5 min

Coverslip 1 drop of Vectashield over each section Apply coverslip Seal with nail polish Store in refrigerator for up to 1 year

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Immunohistochemistry Protocol – Laminin-1

Procedure Directions

Section Preparation 8 µm sections placed on the slide; Allow frozen slides to thaw and dry

Fix Place all slides in acetone - 10 minutes (in freezer) Remove from acetone, allow time to dry, then circle sections with immunopen

Wash & Rehydrate 1x PBS - 5 min, 3 times

Block 10x Blocking Solution (BS): PBS with 10% FBS, 10% GS and 2.5% BSA (1 ml FBS, 1ml GS and 0.25g BSA / 10ml 1X PBS) - 1 hr

Primary Antibody Laminin 111 antibody 1:250 dilution in 1% BS 1 hr

*Don’t Put On Negative Control - Leave block on from last step*

Wash 1% BS - 5 min, 3 times This solution is used for multiple steps and multiple solutions.

Secondary Antibody *In Dark* FITC anti-Rat 1:200 dilution in 1% BS - 60 min place slides in drawer

Wash 1% BS - 5 min DAPI 1:20000 – 5 min 1% BS - 5 min

Coverslip 1 drop of Vectashield over each section Apply coverslip Seal with nail polish Store in refrigerator for up to 1 year

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Immunohistochemistry Protocol – Satellite Cell Content (Pax7/Dystrophin)

Procedure Directions

Section Preparation 8 µm sections placed on the slide; Allow frozen slides to thaw and dry

Fix Place all slides in acetone - 10 minutes (in freezer) Remove from fixative, allow time to dry, then circle sections with immunopen

Wash & Rehydrate 1x PBS - 5 min, 3 times

Block 5% BSA + 10% GS + 0.2% Triton-x 100 - 30 min

Fabs in 5% BSA + 10% GS + 0.2% Triton-x 100 - 30 min 1:20

1st Primary Antibody Pax7 1:2 (Hybridoma Bank, February Lot) in 1% BSA - overnight at 4 *Don’t put on Negative Control - Leave wash on from last step*

Wash 1% BSA (in 1x PBS) - 5 min, 3 times

2nd Primary Antibody Dystrophin (abcam)1:250 dilution in 1% BSA

*Don’t put on Negative Control - Leave wash on from last step*

Wash 1% BSA (in 1x PBS) - 5 min, 3 times

1st Secondary *In Dark* FITC anti-Rabbit 1:400 dilution in 1% BSA - 60 min Antibody

Wash 1% BSA/PBS - 5 min, 3 times

2nd Secondary *In Dark* TRITC anti Mouse 2nd AB Antibody 1 hour (all sections), placed slides in drawer

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Procedure Directions

Wash 1% BSA/PBS—5min

DAPI 1:20000- 5min

1% BSA/PBS—5min

Coverslip 1 drop of Vectashield with DAPI over each section Apply coverslip Seal with nail polish Store in refrigerator for up to 1 year

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Immunohistochemistry Protocol – Satellite Cell Proliferation (Pax7/Ki67)

Procedure Directions

Section Preparation 8 µm sections placed on the slide; Allow frozen slides to thaw and dry

Fix Place all slides in acetone - 10 minutes (in freezer) Remove from acetone, allow time to dry, then circle sections with immunopen

Wash & Rehydrate 1x PBS - 5 min, 3 times

Permeabilization 0.1% Triton X in 1x PBS - 10 min

Wash 1x PBS - 5 min, 3 times

Block 5% BSA (2.5 g/50 ml 1x PBS) - 20 min Fabs in 5% BSA - 30 min 1:20

1st Primary Antibody Ki67 1:100 dilution in 1% BSA - 60 min *Don’t Put On Negative Control - Leave block on from last step*

Wash 1% BSA (in 1x PBS) - 5 min, 3 times 1:5 5% BSA to 1x PBS (12.56 ml 5% BSA/ 50.24 ml 1x PBS) This solution is used for multiple steps and multiple solutions.

2nd Primary Antibody Pax7 1:2 (DSHB, Supernatant) in 1% BSA - overnight at 4 *Don’t put on Negative Control - Leave wash on from last step*

Wash 1% BSA (in 1x PBS) - 5 min, 3 times

1st Secondary *In Dark* FITC anti-Rabbit 1:150 dilution in 1% BSA - 60 min Antibody place slides in drawer

Wash 1% BSA (in 1x PBS) - 5 min, 3 times

2nd Secondary *In Dark* TRITC anti-mouse 1:100 dilution in 1% BSA - 60 min Antibody place slides in drawer

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Procedure Directions

Wash 1% BSA/PBS—5min

DAPI 1:20000- 5min

1% BSA/PBS—5min

Coverslip 1 drop of Vectashield over each section Apply coverslip Seal with nail polish Store in refrigerator for up to 1 year

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Myofiber Cross Sectional Area Measurement Protocol

1. Open Photoshop CSOpen Images For QuantitationUse FITC+DAPI image 2. ImageAdjustmentsBrightness/ContrastsTurn brightness all the way up to 150 (leave contrast alone) 3. AnalysisSet measurement scale (0.31) 4. AnalysisCount tool# all fibers 5. Click Magnetic lasso tooloutline each fiberclick record measurements a. If “Count” column in “record measurements” chart says something other than 1, delete the ones with no Count # 6. Select allExportSave in Fiber Count Images foldername it the slide name 7. The column “Area” is the muscle fiber size data.

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Myonuclear Content Measurement Protocol

Procedure Directions

Capture Images for • Take images at 40x on the inverted fluorescent microscope Assessment • This is approximately 10 to 17 myofibers/image

• Capture 13 FITC/DAPI merged images per animal to be able to assess 200 fibers per animal

Fiber Count Using Adobe Photoshop count tool, manually count the myofibers that have the majority of the fiber within the field of view

Myonuclei Manually count the nuclei that are associated with each pre-numbered myofiber.

*To make the distinction between myonuclei and mononuclear or satellite cells, I only count cells that have a clear dystrophin line on only one side of the cell, and appear to be contained within the myofiber.

Data Compiling Dump all of the fiber count and associated myonuclei data into an excel spreadsheet and take the average of the myonuclei totals for each of the 200 myofibers.

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Muscle Homogenization Protocol

1. Pour liquid N2 in mortar to precool the mortar and pestle 2. Put tissue in mortar and pour liquid N2 over it to keep it cold 3. Crush tissue into fine powder with pestle and put into 2ml eppendorf tube 4. Weight the muscle powder with tube and two empty tubes as control 5. Add 700ul of ice-cold extraction to each tube 6. Vortex and rotate tubes at 4 for 1h 7. Centrifuge tubes at 14000 rpm at 4 for 15min 8. Remove supernatants avoiding the fatty layer on the top 9. Repeat steps 5, 6, 7 adding this time 350ul of ice-cold extraction buffer to each tube. Incubate at 4 for 30min 10. Collect supernatants and mix with the previous one 11. Determine the protein concentrations using a Bradford assay 12. Aliquot samples into 1.5ml microcentrifuge tubes, freeze in liquid N2. Store at -80

Final 5ml 10ml 15ml 20ml 25ml 30ml Hepes 20 mM pH 7.4 100 μl 200μl 300μl 400μl 500μl 600μl EGTA 2 mM 50 100 150 200 250 300 β-glyceropho 50 mM 250 500 750 1000 1250 1500 DTT 1 mM 5 10 15 20 25 30 Na3VO4 1 mM 50 100 150 200 250 300 Triton X-100 1 % 250 500 750 1000 1250 1500 Glycerol 10 % 625 1250 1875 2500 3125 3750 Leupeptin 10 μM 11.9 23.8 35.7 47.6 59.5 71.4 Benzamidine 3mM 50 100 150 200 250 300 Pepstatin A 5 μM 12.5 25 37.5 50 62.5 75 Aprotinin 10 μg/ml 25 50 75 100 125 150 PMSF 1 mM 25 50 75 100 125 150 H2O 3545.6 7091.2 10636.8 14182.4 17728 21273.6

169

Western Blotting Protocol

1. Glass Cassette and Casting Stand Assembly. a. Place the casting frame upright with the pressure cams in the open position and facing forward on flat surface. b. Select a spacer plate of the desired gel thickness and place a short plate on top of it. c. Orient the spacer plate so that the labeling is “up”. Slid the two glass plates into the casting frame, keeping the short plate facing the front of the frame (side with pressure cams). Leaking may occur if the plates are misaligned or oriented incorrectly. d. When the glass plates are in place, engage the pressure cams to secure the glass cassette sandwich in the casting frame. e. Place a comb completely into the assembled gel cassette. Mark the glass plate 1cm below the comb teeth. This is the level to which the resolving gel is poured. Remove the comb. f. Place the casting frame into the casting stand by positioning the casting frame onto the casting gasket while engaging the spring-loaded lever of the casting stand onto the spacer plate. 2. Gel casting. a. Prepare the resolving gel according to the following table. It is useful to add all the reagents except the TEMED to the gel (once the TEMED and APS have been added to the polyacrylamide solution will polymerize in a few minutes)

RUNNING/RESOLVING STACKING 6% 7% 8% 10% 3.5% 4% dd H2O 5.8 11.6 5.5 11.1 5.3 10.6 4.8 9.6 3.85 7.7 3.78 7.5 5 40% Acrylamide 1.5 3 1.75 3.5 2 4 2.5 5 0.43 0.87 0.5 1 1.5 M Tris (pH 8.8) 2.5 5 2.5 5 2.5 5 2.5 5 ------1 M Tris (pH 6.8) ------0.62 1.25 0.62 1.25 10% SDS ul 100 200 100 200 100 200 100 200 50 100 50 100 10% APS ul * 100 200 100 200 100 200 100 200 50 100 50 100 TEMED ul * 10 20 10 20 10 20 10 20 10 20 10 20 10 20 10 20 10 20 10 20 5 10 5 10

b. Pour lower gel solution using disposable plastic pipette or a syringe. Immediately after pouring the gel, gently add a some drops of ddH20 or ethanol on top of the gel to avoid exposure to air. Leave the gel to polymerize (typically 15-20min). A sharp line between water layer and gel indicates completion of polymerization. While waiting for the gel to polymerize start preparing the stacking gel, the samples, marker and running buffer. c. When the polymerization of resolving gel is complete, remove the layer of water down sink, and absorb excess water with whatman paper d. Begin reaction for upper gel again by adding APS and TEMED, swirling gently. Pour the stacking gel directly on top of the resolving gel using a disposable plastic pipette or a syringe. Insert the comb gently and quickly before the gel polymerizes. Leave to polymerize until gel gets firm (generally about 10 min).

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3. Prepare Samples a. Thaw samples on ice b. Determine amount of sample to load. We usually load 50 -100 μg of protein/well in a total volume of 30 μl c. Add water, then 12.5 Laemmli Buffer, then sample to eppendorf tubes (mix them pipetting up and down several times. d. Boil samples 5 min on heat block, and then spin down samples in microcentrifuge to bring all liquid to bottom. e. Samples are now ready to be loaded onto gel.

4. Sample loading and electrophoresis Required materials: - Clean and dry Mini-Protean Tetra cell tank - Electrophoresis module ( Electrode assembly module only for 1 or 2 gels; for 3 or 4 gels also use the companion running module) - Running buffer 700 ml for 2 gels; 1000 ml for 4 gels - If case, buffer dam

4.1 Assembly a. Set the clamping frame to the open position on a clean flat surface. b. Place the first gel cassette (with the short plate facing inward) onto the gel supports. Note that the gel will now rest at 30º angle, tilting away from the center of the clampling frame. Now, place the second gel on the other side of the clamping frame, again by resting the gel onto the supports. If an odd number of gels (1 or 3) is being run, you must use the buffer dam c. Using one hand, gently pull both gels towards each other, making sure they rest firmly and squarely against the green gaskets. d. While gently squeezing the cassettes, keeping constant pressure, slide the green arms of the clamping frame over the gels, locking them into place. e. Fill the assembly with buffer to just under the edge of the outer gel plate. Check if leaks. If not, at this point, the sample wells can be washed-out with running buffer, and sample can be loaded.

4.2 Sample Loading. a. Load samples into each of the assemblies while they are sitting on a flat surface, OUTSIDE of the tank. Load samples slowly to allow them to settle evenly on the bottom of the well 5. TRANSFER 5.1 Preparation for Blotting a. Freeze ice block prior to preparation of blot assembly. Usually is kept at -20º C freeze b. Prepare the transfer buffer. c. Cut membrane and the filter paper to the dimensions of the gel if necessary. d. Remove gel from glass plates (rinse if necessary with running dH2O), cut off upper (stacking) gel and left-hand up corner. Equilibrate gel in transfer buffer for 15 minutes. e. Set up transfer apparatus. • Fill the Criterion Blotter tank with transfer buffer to about 50% of the fill volume.

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• Place a magnetic stir bar inside the tank. (0.8–10 mm) • Place the ice block in the ice block pocket in the back of the cell. Flip down the lever to hold the ice block down. f. Set up the gel/membrane sandwich: 1- Pour chilled transfer buffer into each compartment of the gel/blot assembly tray. 2- Place the membrane (nitrocellulose, PVDF, etc.) in the front/small compartment of the tray. Let it soak while you set up steps f-3–7. 3- Place the cassette in the back/large compartment of the tray: Open the cassette so that the red side with the handle is vertical (anode) and the black side (cathode) is laying horizontal and submerged in transfer buffer. 4- Place a fiber pad on top of the black side of the cassette, submerged in buffer. Push on the fiber pad with gloved finger tips to thoroughly soak the pad. 5- Place a piece of filter paper on top of the fiber pad (it will wet immediately). 6- Gently place the pre-equilibrated gel on top of the filter paper. Use the roller to remove any air bubbles that may be trapped underneath the gel. 7- Take the membrane from the front compartment and place it on top of the gel taking care not to trap any air bubbles. The membrane should not be moved or adjusted once it touches the gel because this can cause data ghost prints and artefacts. If you feel that you must adjust the membrane placement, use a fresh pre-wetted membrane. Use the roller to roll out bubbles. 8- Place a piece of filter paper on top of the membrane. Run the roller gently over the top of the filter paper to remove any air bubbles trapped in the sandwich. 9- Wet a second fiber pad in the front compartment of the tray (where the membrane was soaking) again using finger tips to completely saturate the pad with transfer buffer. Then place the wet fiber pad on top of the second filter paper. 10- Lower the clamp-side of the cassette, and lock in the closed position 5.2 Begin Transfer a. Move the locked cassette into the groove in the blotter tank, aligning the red side of the card with the red electrode. Make sure the magnetic stirrer is free to move.

b. After both cassettes are in place, add the remaining transfer buffer to the fill level marked on the tank. c. Put on the lid, plug the cables into the power supply, and run the blot. The Criterion Blotter has PLATE electrodes so transfer at 100 V during 30 min. If need increase the transfer time be sure that the buffer is not too hot!! d. Upon completion of the run, disassemble the blotting sandwich and remove the membrane for development. Place the membrane into small Tupperware container and rinse briefly 3X with dd H2O. Clean the cell, fiber pads, and cassettes with multiple rinses of deionized water.

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PROCEDURE DETAILS COMMENTS Gels ___4____% stacking _____6__% running

Prepare Add 31mg DTT into 4xLB Samples Boil samples for 5 min Centrifuge 10s Electrophoresis Start at __100V___ Run ___100__v for __30___min Then Run ___200__v for __30___min

Transfer Equilibrate gel/membrane/paper Start at __100___v Transfer ___100__v for __75___min (IN COLD ROOM) Ponceau S staining 1min Rinse membrane with ddH2O 3X Take pictures Cut the membrane between the target bands Rinse membrane with ddH2O 2X

Block 5% BSA + 400ul NaN3 2g BSA powder + 40ml TBST __1___ hour

Primary Antibody Information Antibody Phospho-mTOR (Ser2448) Rabbit Ab (289kda) Phospho-p70s6k (Thr389) Rabbit Ab (70kda) Incubation __overnight__

Rinse 3X Rinse with 0.1%TBS-T, 5min, each

Secondary Antibody Information Antibody __Anti-Rabbit HRP______1:2000_in 2.5% BSA in 0.1% TBST_ Incubation ___1__ hour

Rinse 4X Rinse with 0.1%TBS-T, 10min, each

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