The Role of Toll Like Receptor-4 in Exercise-Induced Myokine Response and Regulation Of

The Role of Toll like Receptor-4 in Exercise-induced Myokine Response and Regulation of

Skeletal Muscle Metabolic Adaptation

Mostafa Mahmoud Abdelaziz Ali

Dissertation submitted to the faculty of the Virginia Polytechnic Institute and State University in partial fulfillment of the requirements for the degree of

Doctor of Philosophy In Human Nutrition, Foods and Exercise

Matthew W. Hulver, Chair Kevin P. Davy Robert W. Grange Madlyn I. Frisard Liwu Li

October 28th, 2016 Blacksburg, VA

Keywords: skeletal muscle, toll-like receptor-4, exercise metabolism

The Role of Toll like Receptor-4 in Exercise-induced Myokine Response and Regulation of Skeletal Muscle Metabolic Adaptation

Mostafa M. Ali ABSTRACT (ACADEMIC) Toll like receptor-4 (TLR4) is a transmembrane inflammatory receptor expressed ubiquitously

on the cell surface of immune cells as well as skeletal muscle and other metabolic tissues. A

compelling body of evidence shows that muscle TLR4 and the downstream cytokine signaling

modulate skeletal muscle metabolism. Intriguingly, skeletal muscle has been demonstrated to

gain favorable inflammatory cytokine-mediated metabolic adaptations in the context of exercise

training. This paradigm suggests a role for muscle TLR4 inflammatory signaling in the

regulation of exercise metabolism. As such, the question arises as to whether exercise stress

response follows similar inflammatory physiological pathways to those activated by other

physical and pathogenic stimuli or not. Therefore, the objective of the present study was to

investigate the role of muscle TLR4 signaling in modulating skeletal muscle cytokine, also

known as myokine, response and metabolic adaptations to exercise. To this end, using Cre-

mediated recombination, we developed a novel muscle-specific TLR4 knockout (mTLR4-/-)

mouse model on C57BL/6JJ background. The differential inflammatory and metabolic responses

between mTLR4-/- mice and wild type (WT) littermates were examined following exposure to either exercise or muscle stimulus. Accordingly, different exercise and muscle contraction modalities were pursued, focusing on voluntary wheel running, forced treadmill training, and in vivo electrical muscle stimulation. Overall, this study introduces a novel muscle-specific TLR4

knockout mouse model and discloses a crucial role for mTLR4 in basal systemic cytokine

homeostasis. Furthermore, our findings identify mTLR4 as a major immunomodulatory effector

of exercise-induced metabolic adaptations and suggest a link between mTLR4 and physiological determinants of maximal aerobic performance.

The Role of Toll like Receptor-4 in Exercise-induced Myokine Response and Regulation of Skeletal Muscle Metabolic Adaptation

Mostafa M. Ali ABSTRACT (PUBLIC) Exercise is an effective health care modality that exerts many physiological and metabolic

benefits. Yet, the magnitude of health outcome differs between individuals, which has

encouraged scientists to study biological factors responsible for variable responses to exercise.

Our bodies respond to exercise as a whole, which requires harmonious communication among

multiple body systems. Skeletal muscle, a major metabolic tissue mainly responsible for bodily movements, plays a key role in whole body energy balance. During exercise skeletal muscle naturally undergoes mechanical and metabolic stress with a subsequent immune reaction known as inflammation. Not only is this exercise-induced inflammatory response known to repair muscle damage, it has also been shown to modulate several of the salutary metabolic effects of exercise. The goal of the study was to better understand how exercise-induced inflammatory response and the subsequent metabolic adjustments are regulated at the level of skeletal muscle.

Our data indicate that mTLR4, an immune receptor imbedded in the surface of muscle cells, modulates the inflammatory signals initiated during exercise. Furthermore, we found that genetically modified mice lacking mTLR4 were unable to develop the normal metabolic adaptations to exercise training. Unlike wild type mice, these mTLR4 deficient mice failed to improve fat and/or glucose utilization after one month of either voluntary wheel running or controlled treadmill training. These findings suggest that defects in this immune receptor, commonly reported with obesity, may alter whole body metabolism and the health outcomes of exercise. Future studies should aim to investigate whether different exercise modalities, such as resistance training, could possibly bypass these limitations induced by mTLR4 abnormities.

ACKNOWLEDGMENTS

First and foremost, I would like to praise and thank Allah (s.w.t) for without Him none of this would have been possible.

I would like to thank my home country, Egypt, for entrusting me the responsibility of learning. I am looking forward to transferring and implementing the knowledge and inspiration I have gained during my PhD study in the United States of America. I was initially inspired by colleagues before me who came to the U.S.A. to seek high quality education. I was impressed by how excited they were about their work and how they got committed to the advancements and betterment of the developing world. As such, I have always wanted to be a part of this intellectual exchange.

I would also like to thank my advisory committee for all their help and guidance over the course of my PhD studies. I would like to express the deepest appreciation to my committee chair

Professor Mathew Hulver who always showed much care and patience with me. He taught me research as well as Metabolic Nutrition, a very special class I will never forget. He also awarded me a precious opportunity where I enjoyed teaching with him for 3 years. This experience has always been a great asset to the development of my academic career. I would like to express my sense of gratitude to Dr. Kevin Davy who was always willing to help me by either sharing knowledge or providing valuable scientific resources. I want to deeply thank Dr. Robert Grange for his constructive feedback and continued scientific guidance throughout my PhD. He always promoted my critical thinking and enforced my analytical skills. Besides, I would also like to offer special thanks to Dr. Madlyn Frisard for her help with my course work. She was always

willing to understand what my challenges were and tried to help me overcome any hitches

hindering my learning process. I would like to recognize Dr. Liwu Li, the expert of

inflammation biology, for his guidance and unlimited generosity. He always had the time and

knowledge to address my questions and provide assistance with his innovative scientific insights.

Besides my advisor committee, I would like to extend my warmest thanks and appreciation to

Dr. Ryan McMillan not only for his insightful comments and encouragement, but also for the

hard questions which incented me to widen my research from various perspectives.

My sincere thanks go to my colleagues in the Hulver Laboratory and in the Integrated Life

Sciences Building (ILSB) who provided me an opportunity to join their team. Without their help

it would not have be possible to conduct this research. It is also a delight to acknowledge my

friends in the Islamic Center of Blacksburg (ICB). In particular, I would like to thank Dr. Sedki

Riad, Ali Nazmi, Hassan Mahsoub, Mohamed Henawy, Hana Alkhalidy, Haithem Taha,

Haitham Elmarakeby, Mohamed Mohamedin, Sulaiman Matarneh, Tariq Abuhamdia, Atia Eisa,

Amr Abed, Ahmed Roman, Ahmed Ghanem, Bassem Mokhtar, Karim said, Abdallah Hassan,

Abdullah Awaysheh, and Abdullah Al-Naib for their friendship and help over the years.

I owe a debt of gratitude to my dear friend Michael Tomaszewicz for giving me a family away

from home and helping me throughout my PhD journey. I would like to convey my heartfelt

thanks to my beloved mother, my brothers Mohamed and Ahmed, and my sisters Fatimah and

Shaymaa for their unflagging love and unconditional support. I would also like to express my

warmest sense of gratitude to my sincere wife Hagar who is such a renewable source of peace and inspiration in my life. Last but certainly not least, I am honored to dedicate this dissertation

to the memory of my beloved father, Mahmoud Abdelaziz Ali, who spared no effort to provide the best possible environment for me to grow up and be who I am. All praise is due to Allah.

TABLE OF CONTENTS

ABSTRACT (ACADEMIC)………………………………………………………………...……ii

ABSTRACT (PUBLIC)………………………………………………………………………….iii

ACKNOWLEDGMENTS………………………………………………………………….…….iv

TABLE OF CONTENTS…………………………………………………………………………vi

LIST OF FIGURES……………………………………………………………………………..viii

LIST OF TABLES………………………………………………………………………………...x

CHAPTER I: INTRODUCTION………………………………………………………………...1

References…………………………………………………………………………………4

CHAPTER II: REVIEW OF LITERATURE……………………………………………………7

Introduction………………………………………………………………………………..7

Exercise as a Stimulus: Emphasis on Skeletal Muscle Cytokine Response………………9

Inflammation and Exercise-induced Muscle Growth ……………………………………11

Inflammation and Exercise-induced Skeletal Muscle Metabolic Response…………..…13

Myokines and Exercise Metabolism……………………………………………………..14

Conclusion……………………………………………………………………………….17

References………………………………………………………………………………..18

CHAPTER III: SPECIFIC AIMS………………………………………………………………24

CHAPTER IV: RESEARCH APPROACH AND EXPRIMENTAL DESIGN…….………….26

Study One: Differential exercise-induced metabolic adaptations between mTLR4-/-mice and WT littermates using voluntary wheel running……………………………………...26

Experimental Design……………………………………………………………..26

Study Two: Differential exercise-induced metabolic adaptations and aerobic capacity between mTLR4-/-mice and WT littermates using treadmill running…………………...27

Experimental Design……………………………………………………………..27

Study Three: Differential exercise-induced serum myokine response between mTLR4-/- mice and WT littermates using acute treadmill running…………………………………28

Experimental Design……………………………………………………………..29

Study Four: Differential muscle contraction-induced myokine gene expression between mTLR4-/-mice and WT littermates using in-vivo electrical muscle stimulation………...29

Experimental Design……………………………………………………………..30

Limitations, Alternative Approaches, and Future Directions……………………………30

Extended Methods……………………………………………………………………….31

CHAPTER V: Muscle-Specific Deletion of Toll-like Receptor-4 Impairs Metabolic Adaptation to Wheel Running in Mice……………………………………………………………………….40

Abstract…………………………………………………………………………………..40

Introduction………………………………………………………………………………41

Material and Methods ……………………………………………………………………42

Results ……………………………………………………………………………………48

Discussion………………………………………………………………………………..57

References………………………………………………………………………………..61

CHAPTER VI: Skeletal Muscle Toll-like Receptor-4 Deficiency Suppresses Exercise-induced Myokine Response and Diminishes Aerobic Training Adaptations in Mice……………………64

Abstract…………………………………………………………………………………..64

Introduction………………………………………………………………………………65

Material and Methods ……………………………………………………………………67

Results ……………………………………………………………………………………73

Discussion………………………………………………………………………………..81

References………………………………………………………………………………..86

CHAPTER VII: Conclusions and Future Directions…………………………………………...89

APPENDICES.………………………………………………………………………………….91

Appendix A: Generation of Skeletal Muscle-Specific TLR4 Knockout Mice.……………...91 vii

LIST OF FIGURES

CHAPTER I

Figure 1. Exercise-induced Myokines and Potential Roles in Modulating Metabolism and Muscle Adaptations………………………………………………………………………………………..2

CHAPTER II

Figure 1. Exercise-inducible Factors Reported to Modulate Physiological and Metabolic Events…………………………………..……………………………………………………….....8

Figure 2. Muscle as an Endocrine Organ………………………………………………………...10

Figure 3. Secondary Phase of Muscle Damage………………………………………………….12

Figure 4. Metabolic Mechanism of IL-6………………………………………………………....16

CHAPTER IV

Figure 1. Study One Timeline: Voluntary Wheel Running……………………………………...27

Figure 2. Study Two Timeline: Chronic Treadmill Training…………………………………....28

Figure 3. Study Three Timeline: Acute Exhaustive Treadmill Running………………………...29

Figure 4. Study Four Timeline: In Vivo Electrical Muscle Stimulation………………………....30

CHAPTER V

Figure 1. mTLR4 protein content is decreased and coupled with lower IRAK4 phosphorylation in mTLR4-/- mice………………………………………………………………………………...49

Figure 2. Wheel running was associated with reductions in body mass and fat mass as well as an increase in fat free mass, however, no genotype differences were observed in wheel running performance……………………………………………………………………………………...50

Figure 3. mTLR4 mediates wheel running-induced increases in substrate oxidation in skeletal muscle……………………………………………………………………………………………52

viii

Figure 4. mTLR4 modulates the oxidative capacity of muscle mitochondrial enzymes in response to wheel running…………………………………………………………………………………55

Figure 5. mTLR4 signaling is required to enrich glucose-6-phosphate metabolite pools in response to wheel running……………………………………………………………………….55

Figure 6. Changes in gene expression of key metabolic and inflammatory regulators in response to wheel running…………………………………………………………………………………56

CHAPTER VI

Figure 1. Schematic representation of the treadmill study design……………………………….68

Figure 2. The effect of the treadmill training protocol on body fat and maximal aerobic capacity…………………………………………………………………………………………..74

Figure 3. Assessment of force production over the course of the in-vivo electrical muscle stimulation protocol……………………………………………………………………………...75

Figure 4. Assessment of inflammatory markers post-exhaustive treadmill running…………….76

Figure 5. Assessment of myokine gene expression in response to in-vivo electrical muscle stimulation………………………………………………………………………………………..77

Figure 6. Assessment of changes in oxidative metabolism in response to treadmill training…...78

Figure 7. Assessment of changes in mitochondrial maximal enzymatic activity in response to treadmill training………………………………………………………………………………....79

Figure 8. Assessment of changes gene expression of key metabolic and inflammatory genes in response to treadmill training………………………………………………………………...…..80

Figure 9. Assessment of changes in GLUT4 protein levels in response to treadmill training…..81

LIST OF TABLES

Table 1. Study One Experimental Design: Voluntary Wheel Running………………………….26

Table 2. Study Two Experimental Design: Chronic Treadmill Training………………………..27

Table 3. Study Three Experimental Design: Acute Exhaustive Treadmill Running ……………28

Table 4. Study Four Experimental Design: In Vivo Electrical Muscle Stimulation ……………29

CHAPTER I

INTRODUCTION

Exercise is a physical stressor that induces a range of physiological responses stimulated by a

variety of molecular signals[1]. The strong association between exercise and the subsequent

elevations in systemic inflammatory markers encouraged researchers to explore the impact of the latter on the regulation of exercise metabolism (Figure 1). In response to acute exercise,

contracting muscles secrete small inflammatory proteins known as cytokines [2]. These muscle-

derived cytokines, recognized as myokines, signal locally in autocrine and paracrine manners as

well as systemically following an endocrine mechanism [3-5]. Circulating myokines, released in response to exercise, are believed to facilitate a crosstalk between active muscle and other body organs such as the liver and adipose tissue [4]. Secretome analysis and mechanistic studies have identified several myokines and their roles in exercise metabolism [6]. Exercise-associated myokines such as interleukin-6 (IL-6), monocyte chemoattractant protein-1 (MCP-1), interleukin-8 (IL-8), and interleukin-15 (IL-15) exhibit varied time course profiles [7, 8] suggesting involvement of different regulatory mechanisms. Several of these myokines, most notably IL-6, are demonstrated to mediate favorable metabolic responses to exercise [9, 10]. In vitro exposure to IL-6 has been shown to stimulate glycogen synthesis and glucose oxidation in human skeletal muscle [11]. It has also been observed that IL-6 infusion in humans augments lipid oxidation with increased reliance on free fatty acids as a source of energy production in skeletal muscle under basal conditions [12].

Figure 1. Exercise-induced Myokines and Potential Roles in Modulating Metabolism and

Muscle Adaptations (Adapted from B So et al. 2014)[13]

Although the release of myokines upon muscle contraction has been known for decades, the underlying mechanisms are still unclear. Several studies proposed different molecular mechanisms regulating these inflammatory cascades triggered by exercise and muscle contraction. Post-exercise muscle glycogen content is reported to be tightly related to levels of circulating cytokines and has been shown to regulate myokine gene expression [14, 15]. Another line of evidence has suggested that exercise-induced intracellular calcium flux as well as production of reactive oxygen species (ROS) activates transcriptional factors promoting myokine synthesis [16-18]. Nonetheless, research undertaken to characterize the mechanisms of exercise- induced myokine release is limited to the effect of changes in the intracellular signaling networks within the active muscle such as an altered metabolite concentration or changes in gene

expression [19]. In fact, skeletal muscle is a part of a dynamic environment rich with potent circulating signaling molecules (e.g. hormones, ROS) that prime muscle metabolism to meet the heightened energy demand during an exercise challenge [20]. Thus, to better understand the interplay between the inflammatory signals and exercise-induced metabolic responses, further research should investigate the role of inflammatory signaling pathways mediated by cell surface receptors that can sense extracellular changes in muscle microenvironment.

Toll-like receptors (TLRs) are cell surface proteins that function in the innate immune system by recognizing microbial pathogens as well as endogenous extracellular signaling molecules [21-

27]. Interestingly, a class of TLRs, primarily TLR4, was found to be highly expressed in skeletal muscle and other major metabolic tissues, suggesting a role in body energy homeostasis [28]. In addition to mediating pathogen-induced cytokine production, a crucial role for TLR4 in modulating skeletal muscle metabolism has been identified [29]. Meanwhile, dysregulation of muscle TLR4 expression was found to generate metabolic perturbation in the context of inflammatory diseases such as obesity and type 2 diabetes [28, 30, 31]. These findings suggest

TLR4 as a potential modulator of exercise metabolism. In concert, similar to a typical response to an endotoxin challenge, a down-regulation in TLR4 gene expression has been observed following prolonged vigorous exercise [32, 33]. Furthermore, prolonged exercise has been documented to stimulate the induction of molecules including heat shock proteins (HSPs) and none esterified fatty acids (NEFAs) that are suggested to be activators of TLR4 [34, 35].

Collectively, these results suggest that TLR4 could be a prime candidate to serve as a central

effector in exercise metabolism [36]. This effect is proposed to work either directly via

modulating key exercise-associated metabolic regulators or indirectly through downstream

myokine-dependent mechanisms. To date however, there is no direct evidence to support this

paradigm. Therefore, the objective of the present study was to investigate the role of muscle

TLR4 signaling in modulating skeletal muscle cytokine response and metabolic adaptations to

exercise. In this study, muscle-specific TLR4 knockout mice and wild type littermates were

utilized. The inflammatory and metabolic responses of each of the genotypes were examined

following exposure to either exercise or muscle stimuli. Accordingly, different exercise and

muscle contraction modalities were pursued, focusing on voluntary wheel running, forced

treadmill training, and in vivo electrical muscle stimulation.

References

1. Radak, Z., et al., Oxygen consumption and usage during physical exercise: the balance between oxidative stress and ROS-dependent adaptive signaling. Antioxid Redox Signal, 2013. 18(10): p. 1208-46. 2. Suzuki, K., et al., Systemic inflammatory response to exhaustive exercise. Cytokine kinetics. Exerc Immunol Rev, 2002. 8: p. 6-48. 3. Pedersen, B.K., Muscles and their myokines. J Exp Biol, 2011. 214(Pt 2): p. 337-46. 4. Schnyder, S. and C. Handschin, Skeletal muscle as an endocrine organ: PGC-1alpha, myokines and exercise. Bone, 2015. 80: p. 115-25. 5. Pedersen, B.K., Edward F. Adolph distinguished lecture: muscle as an endocrine organ: IL-6 and other myokines. J Appl Physiol (1985), 2009. 107(4): p. 1006-14. 6. Scheler, M., et al., Cytokine response of primary human myotubes in an in vitro exercise model. Am J Physiol Cell Physiol, 2013. 305(8): p. C877-86. 7. Louis, E., et al., Time course of proteolytic, cytokine, and myostatin gene expression after acute exercise in human skeletal muscle. J Appl Physiol (1985), 2007. 103(5): p. 1744- 51. 8. Schild, M., et al., Effects of Acute Endurance Exercise on Plasma Protein Profiles of Endurance-Trained and Untrained Individuals over Time. Mediators Inflamm, 2016. 2016: p. 4851935. 9. Smith, L.L., Cytokine hypothesis of overtraining: a physiological adaptation to excessive stress? Med Sci Sports Exerc, 2000. 32(2): p. 317-31. 10. Kelly, M., et al., AMPK activity is diminished in tissues of IL-6 knockout mice: the effect of exercise. Biochem Biophys Res Commun, 2004. 320(2): p. 449-54. 11. Glund, S., et al., Interleukin-6 directly increases glucose metabolism in resting human skeletal muscle. Diabetes, 2007. 56(6): p. 1630-7. 12. Wolsk, E., et al., IL-6 selectively stimulates fat metabolism in human skeletal muscle. Am J Physiol Endocrinol Metab, 2010. 299(5): p. E832-40.

13. So, B., et al., Exercise-induced myokines in health and metabolic diseases. Integrative Medicine Research, 2014. 3(4): p. 172-179. 14. Keller, C., et al., Effect of exercise, training, and glycogen availability on IL-6 receptor expression in human skeletal muscle. J Appl Physiol (1985), 2005. 99(6): p. 2075-9. 15. Nieman, D.C., et al., Post-Exercise Skeletal Muscle Glycogen Related to Plasma Cytokines and Muscle IL-6 Protein Content, but not Muscle Cytokine mRNA Expression. Front Nutr, 2015. 2: p. 27. 16. Fischer, C.P., Interleukin-6 in acute exercise and training: what is the biological relevance? Exerc Immunol Rev, 2006. 12: p. 6-33. 17. Keller, C., et al. Human muscle cells express IL-6 via a Ca2+-dependent pathway. in JOURNAL OF PHYSIOLOGY-LONDON. 2002. CAMBRIDGE UNIV PRESS 40 WEST 20TH ST, NEW YORK, NY 10011-4221 USA. 18. Broholm, C., et al., Exercise induces expression of leukaemia inhibitory factor in human skeletal muscle. J Physiol, 2008. 586(8): p. 2195-201. 19. Peake, J.M., et al., Cytokine expression and secretion by skeletal muscle cells: regulatory mechanisms and exercise effects. Exerc Immunol Rev, 2015. 21: p. 8-25. 20. Melanson, E.L., P.S. MacLean, and J.O. Hill, Exercise improves fat metabolism in muscle but does not increase 24-h fat oxidation. Exerc Sport Sci Rev, 2009. 37(2): p. 93-101. 21. Takeda, K. and S. Akira, Roles of Toll-like receptors in innate immune responses. Genes Cells, 2001. 6(9): p. 733-42. 22. Chen, G.Y. and G. Nunez, Sterile inflammation: sensing and reacting to damage. Nat Rev Immunol, 2010. 10(12): p. 826-37. 23. Kawai, T. and S. Akira, The role of pattern-recognition receptors in innate immunity: update on Toll-like receptors. Nat Immunol, 2010. 11(5): p. 373-84. 24. Ehrchen, J.M., et al., The endogenous Toll-like receptor 4 agonist S100A8/S100A9 (calprotectin) as innate amplifier of infection, autoimmunity, and cancer. J Leukoc Biol, 2009. 86(3): p. 557-66. 25. Bianchi, M.E., HMGB1 loves company. J Leukoc Biol, 2009. 86(3): p. 573-6. 26. Gondokaryono, S.P., et al., The extra domain A of fibronectin stimulates murine mast cells via toll-like receptor 4. J Leukoc Biol, 2007. 82(3): p. 657-65. 27. Miller, Y.I., et al., Toll-like receptor-4 and lipoprotein accumulation in macrophages. Trends Cardiovasc Med, 2009. 19(7): p. 227-32. 28. Jung, U.J. and M.S. Choi, Obesity and its metabolic complications: the role of adipokines and the relationship between obesity, inflammation, insulin resistance, dyslipidemia and nonalcoholic fatty liver disease. Int J Mol Sci, 2014. 15(4): p. 6184-223. 29. Frisard, M.I., et al., Toll-like receptor 4 modulates skeletal muscle substrate metabolism. American Journal of Physiology-Endocrinology and Metabolism, 2010. 298(5): p. E988- E998. 30. Lumeng, C.N., Innate immune activation in obesity. Mol Aspects Med, 2013. 34(1): p. 12-29. 31. McMillan, R.P., et al., Selective overexpression of Toll-like receptor-4 in skeletal muscle impairs metabolic adaptation to high-fat feeding. Am J Physiol Regul Integr Comp Physiol, 2015. 309(3): p. R304-13. 32. Fernandez-Gonzalo, R., et al., TLR4-mediated blunting of inflammatory responses to eccentric exercise in young women. Mediators Inflamm, 2014. 2014: p. 479395.

33. Oliveira, A.G., et al., Physical exercise reduces circulating lipopolysaccharide and TLR4 activation and improves insulin signaling in tissues of DIO rats. Diabetes, 2011. 60(3): p. 784-96. 34. Kaisho, T. and S. Akira, Toll-like receptor function and signaling. Journal of Allergy and Clinical Immunology, 2006. 117(5): p. 979-987. 35. Nguyen, M.A., et al., A subpopulation of macrophages infiltrates hypertrophic adipose tissue and is activated by free fatty acids via Toll-like receptors 2 and 4 and JNK- dependent pathways. Journal of Biological Chemistry, 2007. 282(48): p. 35279-35292. 36. Gleeson, M., et al., The anti-inflammatory effects of exercise: mechanisms and implications for the prevention and treatment of disease. Nature Reviews Immunology, 2011. 11(9): p. 607-615.

CHAPTER II

REVIEW OF LITERATURE

Introduction

Exercise performance is a multifactorial process that impacts multiple body systems in the acute as well as chronic settings [1-4]. Exercise has multiple systemic effects on the major metabolic tissues ranging from skeletal muscle remodeling to improved whole body energetics [5]. Skeletal muscle is the major site of glucose and fatty acids disposal as it accounts for about 40-50% of total body mass [6]. Hence, exercise is recognized as an effective tool to regulate skeletal muscle metabolism and thereby maintain whole body energy homeostasis. Exercise-induced metabolic adjustments are due to the transient nature of multiple exercise inducible factors as well as the accumulation of chronic exercise training effects. Meanwhile, the metabolic events mediated by exercise are quite specific to the intensity, duration, and frequency of a given exercise mode [7].

Exercise stimulates the secretory nature of skeletal muscle to induce marked local and systemic physiological and molecular changes including increased lactate concentration, production of hypoxia inducible factors, changes in hormone levels, synthesis of heat-shock proteins, generation of oxidative stress, and the production of inflammatory cytokines along with many other mediators related to the immune response [8-10]. Among others, these molecular changes are essential to activate cascades of signal transduction modulating skeletal muscle plasticity. It signals in a pleiotropic fashion to promote muscle contractile machinery and to regulate substrate metabolism for optimal performance during an exercise challenge [5]. The transient rises in inflammatory signals play a multidimensional role in acute exercise [11]. Exercising is known to induce acute muscle injury and tissue damage with a subsequent inflammatory response as a physiological mechanism for muscle repair and growth [12]. Furthermore, the role of muscle-

derived inflammatory mediators in eliciting several of the salutary metabolic effects of exercise has received substantial attention in the past two decades [13].

Figure 1. Exercise-inducible Factors Reported to Modulate Physiological and Metabolic

Events

The current literature provides strong evidence supporting a crucial role for inflammatory cytokines in mediating many of the exercise-induced metabolic benefits. Distinguishable from adipocyte-derived inflammation characterized in disease conditions such as obesity, elevations observed in circulating cytokines are released from active muscle upon exposure to exercise stimulus. Moreover, pathological induction of inflammatory cytokines is demonstrated to be a

TLR-mediated event. Toll-like receptor-4 (TLR4) is an inflammatory signaling receptor that is widely expressed in skeletal muscle [14]. Muscle TLR4 is a dual function protein linking inflammation to metabolism as it also modulates carbohydrate and lipid oxidation in basal conditions [15, 16]. On the other hand, obesity and other systemic inflammatory conditions are characterized with muscle TLR4 gene expression abnormalities often accompanied by impaired 8

metabolic events [17, 18]. Surprisingly, research neither aimed at understanding the role of

muscle TLR4 in exercise metabolism nor was it directed towards discerning its putative

contribution to the exercise-induced myokine response. Therefore, the question remains as to whether exercise stress response follows similar inflammatory physiological pathways to those activated by other physical and pathogenic stimuli or not.

This review discusses recent research identifying the major exercise-associated inflammatory

mediators. It also illustrates the role of the inflammatory signals in exercise metabolism and the

mechanisms by which these signals can influence skeletal muscle metabolic adaptations to

exercise. In addition, it proposes possible avenues for further investigations into a possible role

for muscle TLR4 in the regulation of myokine production and metabolic adjustments in response

to exercise.

Exercise as a Stimulus: Emphasis on Skeletal Muscle Cytokine Response

Exercise is a physical stress that modulates several factors, which alter cellular homeostasis and

signaling networks in a variety of tissues. Skeletal muscle is one of the major tissues impacted by

exercise in a multifaceted manner. During exercise, skeletal muscle undergoes morphological

adjustments as well as intracellular changes in gene expression and metabolite and

electrochemical balance. In general, these responses are intrinsic to the active muscle to meet the

energy demand and to sustain contractile function. Furthermore, some of these muscle inherent

signaling networks are not limited to muscle autoregulation, instead it selectively reaches out to

target tissues known to be significant to exercise performance. The dedicated work by Pedersen’s

team and others, has laid a strong foundation for the muscle-mediated immunomodulatory effect of exercise [19-22]. Early in this millennium, a novel concept of skeletal muscle as an endocrine organ has been developed in muscle physiology research [23-26]. Bente Pedersen and colleagues

have theorized how contracting muscle fibers are able to secrete cytokines which they identified

as “myokines” [27]. These muscle secretory factors were reported to carry out endocrine-like

functions in distal tissues and organs, including the adipose tissue [28, 29], colon [30], liver [31],

kidney [32], pancreas, and brain [33]. The mechanistic paradigm of the local and systemic actions of exercise-induced myokines is proposed to work through anti-inflammatory and metabolic pathways [13, 34, 35].

Figure 2. Muscle as an Endocrine Organ(Adapted from BK Pedersen et al. 2012)[36]

Extensive research has been invested in exploring skeletal muscle protein secretory machinery to discover the key exercise-induced myokines and their role in health and disease. To date, at least

12 myokines have been identified. Exercise has been shown to stimulate the production of a class of myokines with paracrine actions on the organ of secretion including leukemia inhibitory factor

(LIF)[37], IL-15 [38], and brain-derived neurotrophic factor (BDNF) [39]. Other myokines reported to further exhibit endocrine functions include the well-established “exercise factor” IL-6

[40], Irisin [28], calprotectin [41], myonectin [31], oncostatin M (OSM) [42] and secreted protein acidic and rich in cysteine (SPARC)[30]. Recent proteome analysis of skeletal muscle- derived myokines detected a marked increase in mRNA and plasma levels of fractalkine

(CX3CL1) and monocyte chemoattractant protein-1(MCP-1) in humans in response to acute endurance exercise [43]. Furthermore, other putative myokines were discovered using the

MyoMouse strategies developed by Kenneth Walsh of Boston University [44]. The MyoMouse is a genetically engineered mouse model expressing a gene (Akt1 gene) that produces proteins that mimic the effects of weight training, such as decreased insulin resistance, blood glucose, and body fat as well as improvements in other metabolic parameters, without an increase in physical activity or a loss of appetite. Using this experimental approach, Walsh and colleagues were able to identify follistatin-like 1 [45] and fibroblast growth factor 21 (FGF21) [46] as contractile- activated myokines.

Inflammation and Exercise-induced Muscle Growth

Unaccustomed exercise induces tissue damage and a subsequent inflammatory response.

Exercise-induced muscle damage progresses into two phases (Figure 3) [47]. The primary damage phase corresponds to the mechanical strain imposed on muscle fibers during exercise, whereas the secondary phase is characterized with the delayed inflammatory response [48, 49].

A compelling body of evidence indicates that inflammatory signals are a key element in exercise-induced muscle remodeling [50]. Clinical studies reported that administering anti- inflammatory drugs (e.g. NSAIDs and Ibuprofen) compromises the post-exercise muscle microenvironment by attenuating protein synthesis and satellite cells proliferation [51, 52].

Furthermore, post-exercise recovery via cold water immersion was shown to diminish strength

outcome and to reduce muscle hypertrophy compared to post-exercise active recovery methods

[53]. As such, a coordinated inflammatory milieu is a necessity for proper tissue repair and restructuring. In this context, muscle-derived cytokines are a class of proteins thought to orchestrate the inflammatory signals that recruit pertinent immune cells and anabolic pathways

[54]. In particular, neutrophils and macrophages are mobilized to the site of injury [55] to

eliminate damaged muscle fragments via ROS- and pro-inflammatory-dependent phagocytic mechanisms [56, 57]. In succession, regeneration processes of traumatized muscle tissue are launched by localized satellite cells [58].

Figure 3. Secondary Phase of Muscle Damage (Adapted from P Baumert et al. 2016) [47]

Exercise type is well-reported as a significant determinant of the magnitude of muscle injury [59,

60]. Consistent data show that eccentric exercise is associated with a greater muscle damage 12

characterized by a higher creatine kinase activity compared with concentric exercise modalities

[61]. Therefore, eccentric, concentric, and other modes of exercise tend to exhibit differential myokine response. Downhill running has been shown to cause a 135% increase in muscle interleukin-1 beta (IL-1 beta) [59], whereas the systemic levels have been reported with a marginal increase [62]. IL-1 beta promotes inflammation by increasing intracellular calcium and nitric oxide (NO), thereby increasing the expression of adhesion molecules, intercellular adhesion molecules (ICAMs) and vascular cell adhesion molecule-1 (VCAM-1) [63, 64].

Similarly, tumor necrosis factor-α (TNF-α) is expressed during the early phase of inflammatory response as its level rises locally in skeletal muscle [65-67] but with subtle systemic elevations following exercise [68]. TNF-α is also implicated in the activation of pro-inflammatory genes and the ubiquitin–proteasome pathway as means to degrade damaged cellular debris [69].

Interestingly, some cytokines such as IL-6 are better described as double inflammatory agents, where they manifest either pro- or anti-inflammatory properties depending on the physiological environment. Downhill running resulted in substantial but delayed elevations (460%) in plasma

IL-6 compared to pre-exercise levels [70]. However, prolonged running is reported to induce the greatest increase in IL-6 concentration in circulation [62, 68, 71, 72]. In vivo and in vitro studies have shown that IL-6 expression plays a significant role in stimulating satellite cells proliferation in response to acute muscle-lengthening contractions [73] and in satellite cell- mediated muscle hypertrophy [74].

Inflammation and Exercise-induced Skeletal Muscle Metabolic Response

Physical exercise imposes immunological and metabolic stress on the body’s organs [75]. As a result, skeletal muscle, which is the major tissue responsible for force generation and movement, undergoes disruptions in the cytoskeleton, fiber organization, and metabolic regulation [62].

Consequently, cellular sensors transmit signal transductions to pair these homeostatic derangements with transcription of inflammatory and metabolically active proteins [76].

Previously, numerous studies were conducted in search of a contractile-inducible molecule/s, an

“exercise factor” that mediates local and systemic metabolism. This led to the discovery of hormone-like cytokine families released from contracting muscles, also known as myokines [77].

These myokines have extensively been implicated in myocellular and global restitution of energy homeostasis following exercise [13]. Nonetheless, muscle is unlikely the only source of these signaling molecules, for instance adipose tissue and liver were reported to secrete metabolically active factors [78-80]. The secretory nature of skeletal muscle and its extracellular structure, equipped with cell surface receptors, produce a two-way communication channel between active muscle and other distant tissues. Herein, we will address the metabolic significance of a key inflammatory family of myokines in exercise metabolism and regulation of skeletal muscle adaptations.

Myokines and Exercise Metabolism

B. K. Pedersen et al. [13] suggested that “cytokines and other peptides that are produced, expressed, and released by muscle fibers and exert either paracrine or endocrine effects should be classified as myokines.” Although the list includes many protein molecules that fulfill the criteria, we will focus only on myokines presenting the utmost contribution to exercise metabolism.

Interleukin-6

Interleukin-6 (IL-6) is a multifunctional cytokine that plays a central role in inflammation, tissue recovery, and metabolism (Figure 4). Its effects are mediated through interaction with its

receptor complex which comprises two membrane proteins, the ligand-binding α-subunit

receptor (IL-6R) and the signal transducing β- subunit, gp130 [81]. The IL-6R exists both in a membrane-bound and a soluble form (sIL-6R). IL-6 and sIL-6R form a binary complex that binds to two molecules of gp130 and leads to IL-6 transduction, which includes activation of

JAK/STAT, ERK, and PI3k signal transduction pathways [82]. Notably, gp130 is expressed ubiquitously on the cell surface, thus IL-6/sIL-6R complex widens the range of cell types that may respond to IL-6 [83].

A great deal of research has been directed to delineate the molecular pathways underlying the beneficial effects of exercise. The massive release of IL-6 from the contracting skeletal muscle has been proposed as one of the molecular signals “a crucial driver” mediating the exercise- induced effects [84]. It has been reported that the release of IL-6 from active muscle is an intensity-dependent event, which is associated with increased muscle glucose uptake and arterial adrenaline concentration. To this end, IL-6 was claimed to be a carbohydrate sensor because of

the inverse relationship between post-exercise muscle glycogen content and plasma IL-6 levels

[85]. In addition, IL-6 was also shown to increase skeletal muscle glycogen synthesis through a

PI3K-dependent mechanism; to replenish the glycogen depot and restore myocellular

homeostasis [86]. Furthermore, several in vitro studies have demonstrated that IL-6 can enhance both skeletal muscles glucose uptake [86, 87] and fat oxidation [86], principally via the activation of AMP-activated protein kinase (AMPK), with the acute but not with long-term

treatment. Conversely, in vivo investigations have demonstrated that irrespective of the mode of

delivery, acutely or chronically, IL-6 administered to rats for 14 days improved basal insulin sensitivity and promoted glucose disposal. Moreover, while the IL-6 intervention increased circulating fatty acids, it did not increase lipid accumulation in either skeletal muscle or the liver

which was found to be coupled with increased muscle protein content of both PPAR-α and UCP2

[88]. In accordance, IL-6 knockout mice develop late onset obesity and impaired glucose tolerance [89]. Interestingly, IL-6 offers further metabolic merits through anti-inflammatory

mechanisms. IL-6 stimulates the production of anti-inflammatory cytokines and suppression of

TNF-α, which has been reported to have a role in insulin resistance in humans [90]. As such

muscle-derived IL-6 is likely to protect against TNF- α -induced insulin resistance as well as

exercise-induced muscle damage.

Figure 4. Metabolic Mechanism of IL-6 (Adapted from BK Pedersen et al. 2008) [72]

On the other hand, it has been shown that insulin responsive tissues may respond differently to

IL-6. While skeletal muscle appears to be the major target for the reported beneficial effects of

IL-6, Weigert et al. [91] confirmed that, in the liver of mice, IL-6 activates two inhibitory mechanisms that are implicated in insulin resistance, marked induction of SOCS-3 and

phosphorylation of IRS-1. Intriguingly, IL-6 had no substantial role in these insulin desensitization mechanisms in skeletal muscle. Furthermore, skeletal muscle and adipose tissue show differential responses to IL-6 stimulation. The infusion of recombinant human IL-6 (rhIL-

6) was documented to activate skeletal muscle lipolysis via STAT3-dependent mechanisms, whereas this stimulatory effect was absent in adipose tissue. These results suggest that a transient increase in IL-6 within physiological levels selectively promote fat catabolism in skeletal muscle, but not in adipose tissue [92].

In summary, although it is evident that IL-6 mediates exercise-modulated muscle metabolism,

different variables should be considered while interpreting research data. IL-6 is a

multifunctioning myokine with pro- and anti-inflammatory properties. As such, the action of IL-

6 is specific to tissue type, IL-6 levels, time course of IL-6 exposure, causes and components of

inflammatory environment. Another important point not to be overlooked in animal studies is the

fact that mouse and human IL-6 share only about 42% sequence identity [93]. Thus, data

discrepancies might be observed in human trials compared to mouse studies. These confounding

factors, in turn, may give rise to the controversy of the role of IL-6 in modulating substrate

metabolism and energy homeostasis. Therefore, the signaling potential of IL-6 in whole body

physiology is incompletely understood, so research is still needed to model the IL-6 role

considering the aforementioned determinants.

Conclusion

Exercise has a great therapeutic value for preventing and treating modern epidemics of obesity,

type II diabetes, cardiovascular complications, and likely many other diseases. Exercise

metabolism is a major entry point where systemic control of energy homeostasis is transferred

from skeletal muscle to other body organs. The cytokine secretory capacity of skeletal muscle

transformed our understanding of the complex relationship between inflammation and exercise

metabolism. Apparently a healthy inflammatory profile is a target in quest for flexible

inflammatory and metabolic responses to exercise. That being said, differences in exercise-

induced cytokine response in healthy individuals and others with inflammatory conditions (e.g.

aged, obese) after exercise have not yet been characterized. Further research should study

whether the altered inflammatory profile of some disease states (e.g. obesity, type II diabetes,

and aging) could induce metabolic limitations attenuating exercise-concomitant benefits.

References

1. Navarro, A., et al., Beneficial effects of moderate exercise on mice aging: survival, behavior, oxidative stress, and mitochondrial electron transfer. Am J Physiol Regul Integr Comp Physiol, 2004. 286(3): p. R505-11. 2. Chen, C.H., et al., Benefits of exercise training and the correlation between aerobic capacity and functional outcomes and quality of life in elderly patients with coronary artery disease. Kaohsiung J Med Sci, 2014. 30(10): p. 521-30. 3. Yamamoto, S., et al., Effects of resistance training on muscle strength, exercise capacity, and mobility in middle-aged and elderly patients with coronary artery disease: A meta- analysis. J Cardiol, 2016. 68(2): p. 125-34. 4. Colcombe, S.J., et al., Aerobic fitness reduces brain tissue loss in aging humans. J Gerontol A Biol Sci Med Sci, 2003. 58(2): p. 176-80. 5. Ferraro, E., et al., Exercise-induced skeletal muscle remodeling and metabolic adaptation: redox signaling and role of autophagy. Antioxid Redox Signal, 2014. 21(1): p. 154-76. 6. Nader, G.A., Molecular determinants of skeletal muscle mass: getting the "AKT" together. Int J Biochem Cell Biol, 2005. 37(10): p. 1985-96. 7. Haas, T.L., et al., Exercise training and peripheral arterial disease. Compr Physiol, 2012. 2(4): p. 2933-3017. 8. Radak, Z., et al., Oxygen consumption and usage during physical exercise: the balance between oxidative stress and ROS-dependent adaptive signaling. Antioxid Redox Signal, 2013. 18(10): p. 1208-46. 9. Hartwig, S., et al., Secretome profiling of primary human skeletal muscle cells. Biochim Biophys Acta, 2014. 1844(5): p. 1011-7. 10. Scheler, M., et al., Cytokine response of primary human myotubes in an in vitro exercise model. Am J Physiol Cell Physiol, 2013. 305(8): p. C877-86. 11. Covarrubias, A.J. and T. Horng, IL-6 strikes a balance in metabolic inflammation. Cell Metab, 2014. 19(6): p. 898-9. 18

12. Nunes-Silva, A., et al., Treadmill exercise induces neutrophil recruitment into muscle tissue in a reactive oxygen species-dependent manner. An intravital microscopy study. PLoS One, 2014. 9(5): p. e96464. 13. Pedersen, B.K., et al., Role of myokines in exercise and metabolism. J Appl Physiol (1985), 2007. 103(3): p. 1093-8. 14. Vaure, C. and Y. Liu, A comparative review of toll-like receptor 4 expression and functionality in different animal species. Front Immunol, 2014. 5: p. 316. 15. Frisard, M.I., et al., Toll-like receptor 4 modulates skeletal muscle substrate metabolism. American Journal of Physiology-Endocrinology and Metabolism, 2010. 298(5): p. E988- E998. 16. Kim, J.J. and D.D. Sears, TLR4 and Insulin Resistance. Gastroenterol Res Pract, 2010. 2010. 17. Schenk, S., M. Saberi, and J.M. Olefsky, Insulin sensitivity: modulation by nutrients and inflammation. J Clin Invest, 2008. 118(9): p. 2992-3002. 18. Reyna, S.M., et al., Elevated toll-like receptor 4 expression and signaling in muscle from insulin-resistant subjects. Diabetes, 2008. 57(10): p. 2595-602. 19. Goh, J., N. Niksirat, and K.L. Campbell, Exercise training and immune crosstalk in breast cancer microenvironment: exploring the paradigms of exercise-induced immune modulation and exercise-induced myokines. Am J Transl Res, 2014. 6(5): p. 422-38. 20. Pedersen, B.K., et al., Exercise-induced immunomodulation--possible roles of neuroendocrine and metabolic factors. Int J Sports Med, 1997. 18 Suppl 1: p. S2-7. 21. Pedersen, B.K. and A.D. Toft, Effects of exercise on lymphocytes and cytokines. Br J Sports Med, 2000. 34(4): p. 246-51. 22. Lancaster, G.I. and M.A. Febbraio, The immunomodulating role of exercise in metabolic disease. Trends Immunol, 2014. 35(6): p. 262-9. 23. Pedersen, B.K., Exercise-induced myokines and their role in chronic diseases. Brain Behav Immun, 2011. 25(5): p. 811-6. 24. Roca-Rivada, A., et al., Muscle tissue as an endocrine organ: comparative secretome profiling of slow-oxidative and fast-glycolytic rat muscle explants and its variation with exercise. J Proteomics, 2012. 75(17): p. 5414-25. 25. Schnyder, S. and C. Handschin, Skeletal muscle as an endocrine organ: PGC-1alpha, myokines and exercise. Bone, 2015. 80: p. 115-25. 26. Steensberg, A., et al., Production of interleukin-6 in contracting human skeletal muscles can account for the exercise-induced increase in plasma interleukin-6. J Physiol, 2000. 529 Pt 1: p. 237-42. 27. Ostrowski, K., et al., Pro- and anti-inflammatory cytokine balance in strenuous exercise in humans. J Physiol, 1999. 515 ( Pt 1): p. 287-91. 28. Bostrom, P., et al., A PGC1-alpha-dependent myokine that drives brown-fat-like development of white fat and thermogenesis. Nature, 2012. 481(7382): p. 463-8. 29. Quinn, L.S., et al., Oversecretion of interleukin-15 from skeletal muscle reduces adiposity. Am J Physiol Endocrinol Metab, 2009. 296(1): p. E191-202. 30. Aoi, W., et al., A novel myokine, secreted protein acidic and rich in cysteine (SPARC), suppresses colon tumorigenesis via regular exercise. Gut, 2013. 62(6): p. 882-9. 31. Seldin, M.M., et al., Myonectin (CTRP15), a novel myokine that links skeletal muscle to systemic lipid homeostasis. J Biol Chem, 2012. 287(15): p. 11968-80.

32. Ebert, T. and S. Kralisch, Newly discovered myokines in chronic kidney disease. Pol Arch Med Wewn, 2016. 126(7-8): p. 457-8. 33. Engler, D., Hypothesis: Musculin is a hormone secreted by skeletal muscle, the body's largest endocrine organ. Evidence for actions on the endocrine pancreas to restrain the beta-cell mass and to inhibit insulin secretion and on the hypothalamus to co-ordinate the neuroendocrine and appetite responses to exercise. Acta Biomed, 2007. 78 Suppl 1: p. 156-206. 34. Steensberg, A., et al., IL-6 enhances plasma IL-1ra, IL-10, and cortisol in humans. Am J Physiol Endocrinol Metab, 2003. 285(2): p. E433-7. 35. Kawanishi, N., et al., Exercise training inhibits inflammation in adipose tissue via both suppression of macrophage infiltration and acceleration of phenotypic switching from M1 to M2 macrophages in high-fat-diet-induced obese mice. Exerc Immunol Rev, 2010. 16: p. 105-18. 36. Pedersen, B.K. and M.A. Febbraio, Muscles, exercise and obesity: skeletal muscle as a secretory organ. Nat Rev Endocrinol, 2012. 8(8): p. 457-65. 37. Broholm, C., et al., Exercise induces expression of leukaemia inhibitory factor in human skeletal muscle. J Physiol, 2008. 586(8): p. 2195-201. 38. Ostrowski, K., et al., A trauma-like elevation of plasma cytokines in humans in response to treadmill running. J Physiol, 1998. 513 ( Pt 3): p. 889-94. 39. Matthews, V.B., et al., Brain-derived neurotrophic factor is produced by skeletal muscle cells in response to contraction and enhances fat oxidation via activation of AMP- activated protein kinase. Diabetologia, 2009. 52(7): p. 1409-18. 40. Pedersen, B.K., et al., Exercise and cytokines with particular focus on muscle-derived IL- 6. Exerc Immunol Rev, 2001. 7: p. 18-31. 41. Mortensen, O.H., et al., Calprotectin is released from human skeletal muscle tissue during exercise. J Physiol, 2008. 586(14): p. 3551-62. 42. Hojman, P., et al., Exercise-induced muscle-derived cytokines inhibit mammary cancer cell growth. Am J Physiol Endocrinol Metab, 2011. 301(3): p. E504-10. 43. Catoire, M., et al., Identification of human exercise-induced myokines using secretome analysis. Physiol Genomics, 2014. 46(7): p. 256-67. 44. Walsh, K., Adipokines, myokines and cardiovascular disease. Circ J, 2009. 73(1): p. 13- 8. 45. Ouchi, N., et al., Follistatin-like 1, a secreted muscle protein, promotes endothelial cell function and revascularization in ischemic tissue through a nitric-oxide synthase- dependent mechanism. J Biol Chem, 2008. 283(47): p. 32802-11. 46. Izumiya, Y., et al., FGF21 is an Akt-regulated myokine. FEBS Lett, 2008. 582(27): p. 3805-10. 47. Baumert, P., et al., Genetic variation and exercise-induced muscle damage: implications for athletic performance, injury and ageing. Eur J Appl Physiol, 2016. 116(9): p. 1595- 625. 48. Howatson, G. and K.A. van Someren, The prevention and treatment of exercise-induced muscle damage. Sports Med, 2008. 38(6): p. 483-503. 49. Schoenfeld, B.J., The mechanisms of muscle hypertrophy and their application to resistance training. J Strength Cond Res, 2010. 24(10): p. 2857-72. 50. Tidball, J.G., Inflammatory processes in muscle injury and repair. Am J Physiol Regul Integr Comp Physiol, 2005. 288(2): p. R345-53.

51. Trappe, T.A., et al., Effect of ibuprofen and acetaminophen on postexercise muscle protein synthesis. Am J Physiol Endocrinol Metab, 2002. 282(3): p. E551-6. 52. Mikkelsen, U.R., et al., Local NSAID infusion inhibits satellite cell proliferation in human skeletal muscle after eccentric exercise. J Appl Physiol (1985), 2009. 107(5): p. 1600-11. 53. Roberts, L.A., et al., Post-exercise cold water immersion attenuates acute anabolic signalling and long-term adaptations in muscle to strength training. J Physiol, 2015. 593(18): p. 4285-301. 54. Smith, L.L., et al., Cytokines and cell adhesion molecules associated with high-intensity eccentric exercise. Eur J Appl Physiol, 2000. 82(1-2): p. 61-7. 55. Beaton, L.J., M.A. Tarnopolsky, and S.M. Phillips, Contraction-induced muscle damage in humans following calcium channel blocker administration. J Physiol, 2002. 544(Pt 3): p. 849-59. 56. Cannon, J.G., et al., Increased interleukin 1 beta in human skeletal muscle after exercise. Am J Physiol, 1989. 257(2 Pt 2): p. R451-5. 57. Nguyen, H.X. and J.G. Tidball, Interactions between neutrophils and macrophages promote macrophage killing of rat muscle cells in vitro. J Physiol, 2003. 547(Pt 1): p. 125-32. 58. Tidball, J.G., Mechanisms of muscle injury, repair, and regeneration. Compr Physiol, 2011. 1(4): p. 2029-62. 59. Fielding, R.A., et al., Acute phase response in exercise. III. Neutrophil and IL-1 beta accumulation in skeletal muscle. Am J Physiol, 1993. 265(1 Pt 2): p. R166-72. 60. Nosaka, K. and M. Newton, Concentric or eccentric training effect on eccentric exercise- induced muscle damage. Med Sci Sports Exerc, 2002. 34(1): p. 63-9. 61. Pokora, I., et al., Effects of downhill and uphill exercises of equivalent submaximal intensities on selected blood cytokine levels and blood creatine kinase activity. Biol Sport, 2014. 31(3): p. 173-8. 62. Peake, J., K. Nosaka, and K. Suzuki, Characterization of inflammatory responses to eccentric exercise in humans. Exerc Immunol Rev, 2005. 11: p. 64-85. 63. Burgazli, K.M., et al., Importance of large conductance calcium-activated potassium channels (BKCa) in interleukin-1b-induced adhesion of monocytes to endothelial cells. Eur Rev Med Pharmacol Sci, 2014. 18(5): p. 646-56. 64. Dinarello, C.A., Immunological and inflammatory functions of the interleukin-1 family. Annu Rev Immunol, 2009. 27: p. 519-50. 65. Peake, J.M., et al., Cytokine expression and secretion by skeletal muscle cells: regulatory mechanisms and exercise effects. Exerc Immunol Rev, 2015. 21: p. 8-25. 66. Hamada, K., et al., Senescence of human skeletal muscle impairs the local inflammatory cytokine response to acute eccentric exercise. FASEB J, 2005. 19(2): p. 264-6. 67. Cannon, J.G. and B.A. St Pierre, Cytokines in exertion-induced skeletal muscle injury. Mol Cell Biochem, 1998. 179(1-2): p. 159-67. 68. Suzuki, K., et al., Systemic inflammatory response to exhaustive exercise. Cytokine kinetics. Exerc Immunol Rev, 2002. 8: p. 6-48. 69. Reid, M.B. and Y.P. Li, Tumor necrosis factor-alpha and muscle wasting: a cellular perspective. Respir Res, 2001. 2(5): p. 269-72. 70. Peake, J.M., et al., Exercise-induced muscle damage, plasma cytokines, and markers of neutrophil activation. Med Sci Sports Exerc, 2005. 37(5): p. 737-45.

71. Reihmane, D., et al., Increase in IL-6, TNF-alpha, and MMP-9, but not sICAM-1, concentrations depends on exercise duration. Eur J Appl Physiol, 2013. 113(4): p. 851-8. 72. Pedersen, B.K. and M.A. Febbraio, Muscle as an endocrine organ: focus on muscle- derived interleukin-6. Physiol Rev, 2008. 88(4): p. 1379-406. 73. McKay, B.R., et al., Association of interleukin-6 signalling with the muscle stem cell response following muscle-lengthening contractions in humans. PLoS One, 2009. 4(6): p. e6027. 74. Serrano, A.L., et al., Interleukin-6 is an essential regulator of satellite cell-mediated skeletal muscle hypertrophy. Cell Metab, 2008. 7(1): p. 33-44. 75. Pedersen, B.K. and L. Hoffman-Goetz, Exercise and the immune system: regulation, integration, and adaptation. Physiol Rev, 2000. 80(3): p. 1055-81. 76. Egan, B. and J.R. Zierath, Exercise metabolism and the molecular regulation of skeletal muscle adaptation. Cell Metab, 2013. 17(2): p. 162-84. 77. Pedersen, B.K., et al., Searching for the exercise factor: is IL-6 a candidate? J Muscle Res Cell Motil, 2003. 24(2-3): p. 113-9. 78. Golbidi, S. and I. Laher, Exercise induced adipokine changes and the metabolic syndrome. J Diabetes Res, 2014. 2014: p. 726861. 79. Velloso, C.P., Regulation of muscle mass by growth hormone and IGF-I. Br J Pharmacol, 2008. 154(3): p. 557-68. 80. Yakar, S., et al., Liver-specific igf-1 gene deletion leads to muscle insulin insensitivity. Diabetes, 2001. 50(5): p. 1110-8. 81. Hammacher, A., R.J. Simpson, and E.C. Nice, The interleukin-6 (IL-6) partial antagonist (Q159E,T162P)IL-6 interacts with the IL-6 receptor and gp130 but fails to induce a stable hexameric receptor complex. J Biol Chem, 1996. 271(10): p. 5464-73. 82. Kojima, H., et al., IL-6-STAT3 signaling and premature senescence. JAKSTAT, 2013. 2(4): p. e25763. 83. Taga, T. and T. Kishimoto, Gp130 and the interleukin-6 family of cytokines. Annu Rev Immunol, 1997. 15: p. 797-819. 84. Petersen, A.M. and B.K. Pedersen, The anti-inflammatory effect of exercise. J Appl Physiol, 2005. 98(4): p. 1154-62. 85. Helge, J.W., et al., The effect of graded exercise on IL-6 release and glucose uptake in human skeletal muscle. J Physiol, 2003. 546(Pt 1): p. 299-305. 86. Al-Khalili, L., et al., Signaling specificity of interleukin-6 action on glucose and lipid metabolism in skeletal muscle. Mol Endocrinol, 2006. 20(12): p. 3364-75. 87. Glund, S., et al., Interleukin-6 directly increases glucose metabolism in resting human skeletal muscle. Diabetes, 2007. 56(6): p. 1630-7. 88. Holmes, A.G., et al., Prolonged interleukin-6 administration enhances glucose tolerance and increases skeletal muscle PPARalpha and UCP2 expression in rats. J Endocrinol, 2008. 198(2): p. 367-74. 89. Wallenius, V., et al., Interleukin-6-deficient mice develop mature-onset obesity. Nat Med, 2002. 8(1): p. 75-9. 90. Plomgaard, P., et al., Tumor necrosis factor-alpha induces skeletal muscle insulin resistance in healthy human subjects via inhibition of Akt substrate 160 phosphorylation. Diabetes, 2005. 54(10): p. 2939-45.

91. Weigert, C., et al., Direct cross-talk of interleukin-6 and insulin signal transduction via insulin receptor substrate-1 in skeletal muscle cells. J Biol Chem, 2006. 281(11): p. 7060-7. 92. Wolsk, E., et al., IL-6 selectively stimulates fat metabolism in human skeletal muscle. Am J Physiol Endocrinol Metab, 2010. 299(5): p. E832-40. 93. Pedersen, B.K. and M.A. Febbraio, Point: Interleukin-6 does have a beneficial role in insulin sensitivity and glucose homeostasis. J Appl Physiol, 2007. 102(2): p. 814-6.

CHAPTER III

SPECIFIC AIMS

Overview: The overall goal of this study is to investigate the significance of the skeletal muscle

TLR4 (mTLR4) in modulating exercise-induced metabolic adaptations as well as post-exercise myokine production and how mTLR4 deletion influences endurance performance in mice.

Towards this goal, we developed a muscle-specific TLR4 knockout mouse (mTLR4-/-) on

C57BL/6J background and utilized different exercise and in-vivo muscle contraction modalities.

Working Hypothesis: We hypothesize that mTLR4 mediates key myokine responses and metabolic adaptations to exercise; and that the lack of its inflammatory signals limits endurance performance.

Specific Aim 1: To determine the role of mTLR4 as a modulator of exercise-induced metabolic

adaptations

Objective 1: To examine the interaction between voluntary exercise training and the muscle-

specific TLR4 deletion and their effect on the metabolic adaptations in mTLR4-/- mice versus

wild type (WT) littermates

Specific Aim 2: To determine the role of mTLR4 as a determinant of exercise capacity

Objective 2: To examine the interaction between standardized exercise training and the muscle- specific TLR4 deletion and their effect on the exercise capacity and metabolic adaptations in mTLR4-/- mice versus WT littermates

Specific Aim 3: To determine the role of mTLR4 as a mediator of myokine production in

response to exercise and muscle contraction 24

Objective 3: To examine the interaction between the muscle-specific TLR4 deletion and an acute bout of exercise and their effect on serum cytokine levels in mTLR4-/- mice versus WT littermates

Objective 4: To examine the interaction between the muscle-specific TLR4 deletion and

electrical muscle stimulation (EMS) and their effect on myokine gene expression in TLR4-/- mice

versus WT littermates

CHAPTER IV

RESEARCH APPROACH AND EXPRIMENTAL DESIGN

Study One: Differential exercise-induced metabolic adaptations between mTLR4-/-mice and WT littermates using voluntary wheel running

Table 1. Study One Experimental Design: Voluntary Wheel Running

Genotype WT mTLR4-/-

Exercise Condition Sedentary Wheel Running Sedentary Wheel (n=7) (n=7) (n=7) Running (n=7)

Experimental Design.

Voluntary wheel running will be used where 28 mTLR4-/- and WT mice will be randomized to either a sedentary control or exercise group with 24 hour-free access to the running wheels,

Table 1. Body composition parameters will be assessed prior to and after the completion of the exercise program. Mice will be maintained on the wheel running protocol for 4 weeks. Mice will be sacrificed after 24 hours of the last exposure to the wheels. Blood samples will be collected using cardiac puncture. Skeletal muscle (gastrocnemius, quadriceps, and soleus) will be harvested for substrate oxidation (fatty acid, pyruvate, glucose) and maximal enzymatic activity assays, mitochondria bioenergetics, and gene expression analysis of key metabolic and inflammatory markers.

Figure 1. Study One Timeline: Voluntary Wheel Running

Study Two: Differential exercise-induced metabolic adaptations and aerobic capacity between mTLR4-/-mice and WT littermates using treadmill running

Table 2. Study Two Experimental Design: Chronic Treadmill Training

Genotype WT mTLR4-/-

Exercise Condition Sedentary Treadmill Sedentary Treadmill (n=8) Training (n=8) (n=8) Training (n=8)

Experimental Design.

Forced treadmill running will be used where 32 mTLR4-/- and WT mice will be randomized to either a sedentary control or exercise group, Table 2. Mice will be acclimated to the treadmill for

2 days. Body composition parameters and maximal aerobic capacity will be assessed prior to and after the completion of the exercise program. Furthermore, mice will perform a moderate intensity 50-minute run 5 days/week for 4 weeks. The protocol will progress into two phases.

The first two weeks mice will run at 0.30 m/sec where the second two weeks will be at 0.33

m/sec. Mice will be sacrificed 36 hours after the last bout of exercise. Skeletal muscle

(gastrocnemius, quadriceps, and soleus) will be harvested for substrate oxidation (fatty acid,

pyruvate, glucose) and maximal enzymatic activity assays, mitochondria bioenergetics, and gene

expression analysis of key metabolic (e.g. PGC1-α, PPARr-Delta) and inflammatory markers

(IL-6, MCP-1).

Figure 2. Study Two Timeline: Chronic Treadmill Training

Study Three: Differential exercise-induced serum myokine response between mTLR4-/- mice and WT littermates using acute treadmill running

Table 3. Study Three Experimental Design: Acute Exhaustive Treadmill Running

Genotype WT mTLR4-/-

Condition (Time point) Pre-exercise Post-exercise Pre-exercise Post-exercise serum sample serum sample serum sample serum sample (n=6) (n=6) (n=6) (n=6)

Experimental Design.

Forced acute treadmill running will be used and 12 mTLR4-/- and wild type mice will undergo an acute bout of prolonged endurance exercise where mice will run until exhaustion at 0.3/sec,

table 3. Blood samples will be drawn from the saphenous vein prior to and immediately after the

exercise bout. Serum cytokine profile will be analyzed to monitor levels of exercise-induced

inflammatory markers (e.g. IL-6, MCP-1).

Figure 3. Study Three Timeline: Acute Exhaustive Treadmill Running

Study Four: Differential muscle contraction-induced myokine gene expression between mTLR4-/-mice and WT littermates using in-vivo electrical muscle stimulation

Table 4. Study Four Experimental Design: In Vivo Electrical Muscle Stimulation

Genotype WT mTLR4-/-

Condition In vivo EMS Control In vivo EMS Control (n=9) (n=9) (n=9) (n=9)

Experimental Design.

In this study 18 mTLR4-/- and wild type mice will undergo in vivo electrical muscle stimulation,

table 4. Every mouse from each group will serve as both an experimental and control unit

simultaneously. One leg will undergo in vivo electrical stimulation to induce muscle contraction

whilst the other leg will serve as a non-stimulated control. Mice will be sacrificed 3 hours post-

stimulation. Skeletal muscle (gastrocnemius, quadriceps, and soleus) will be harvested for gene

expression assays to quantify mRNA level of key inflammatory myokines (IL-6, MCP-1).

Figure 4. Study Four Timeline: In Vivo Electrical Muscle Stimulation

Limitations, Alternative Approaches, and Future Directions:

The major limitation is the breeding rate of the mTLR4-/- mouse model, since it is domestic to

our laboratory and is not commercially available. This breeding concern could hinder the ability

to obtain a sample size that enables the statistical power sufficient to detect significant (P<0.05)

differences between different experimental groups. Furthermore, mTLR4 deletion selectively

knockout muscle TLR4 gene expression, however, many other immune and fat cells are inherent

to muscle microenvironment. Contamination with non-muscle cells will likely reveal

confounding results with false positive inflammatory gene expression. A proposed alternative

approach to study the role of TLR4 in skeletal muscle during exercise is using C2C12 skeletal

myotubes and gene silencing techniques to obtain dysfunctional mTLR4. As well, isolating

satellite cells from mTLR4-/- mice will provide high quality pure knockout muscle cells.

Furthermore, electrical pulse stimulation for cultured myocytes has become a well-established

technique that mimics exercise-induced muscle contraction. This study will lead to a better

understating of the interplay between inflammatory signals and metabolic adaptations to exercise

and whether the altered inflammatory profile some disease states (e.g. obesity, type II diabetes)

could induce metabolic limitations attenuating concomitant benefits of exercise as a health tool.

Future studies could aim at investigating the discrepancies in the metabolic adaptations seen

potentially among lean and obese individuals in response to exercise. Furthermore, future studies

could help better understand the exercise-drug interactions as related to the relevance of administering, or lack thereof, anti-inflammatory therapies during an exercise regimen.

Extended Methods

Animal husbandry and generation of mTLR4-/- mice. Animal studies will be performed

under an approved protocol by the Institutional Animal Care and Use Committee (IACUC) at

Virginia Tech. The muscle-specific TLR4 knockout (mTLR4-/-) mouse model on C57BL/6J

background was created at the Transgenic Core at the Pennington Biomedical Research Center

using the Cre-mediated recombination driven by the muscle creatine kinase promoter (Appendix

A). Male mTLR4-/- and wild type (WT) littermates will be maintained on a 12-hour light/dark

cycle and fed standard chow diet. All experiments will be performed following an overnight fast

(10-12 hours), unless otherwise stated. To confirm muscle-specific deletion of TLR4, animals

will be sacrificed and flexor digitorum brevis (FDB) muscle will be isolated. The FDB muscle

group will be used because isolation of single muscle fibers limits confounding influences of

non-muscle cell types that are present in whole muscle groups (e.g. gastrocnemius and quadriceps).

Western blot analysis. Western blot analysis will be performed using skeletal muscle

homogenates prepared in 50 mM Hepes, (pH7.5), 15 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 2

mM EDTA, 10% glycerol, 1 % Triton x-100, 10 mM NaP2O7, 100 mM NaFl, 10 mM PMSF,

and 10 g/mL aprotinin. Proteins (~30 µg per sample) will be separated using a 10% Criterion-

Tris-HCl gel (Bio-Rad, Hercules, CA) and subsequently transferred to PVDF membrane (Bio-

Rad, Hercules, CA). PVDF membrane will be probed with primary antibody against genes of

interest.

FDB muscle fibers isolation and preparation. Mice will be euthanized using CO2

asphyxiation followed by cervical dislocation according to Virginia Tech IACUC protocols. The

hind foot will be cut off above the ankle joint. The foot will be pinned with the plantar side

facing up onto a flat foam board. Skin will be reflected starting from the proximal side of the

foot, at the ankle, and will be pulled upwards toward the toes. The FDB muscle bundle will be

dissected by cutting the tendon at the heel and pulling it upward while detaching the muscle

fibers from other adjacent tissues. The FDB muscle bundle will be removed by cutting at the

insertion tendons attached to each of the four lateral toes. The FDB muscle will be immediately

placed in a tube with 500 µL of dissociation buffer containing cell lysis buffer mixed with

protease and phosphatase inhibitors (1/100) as well as collagenase (4 mg/mL). The tube will be

rocked in a horizontal position on a shaker for approximately 90 minutes at 37o C. Digested

muscle will be transferred into a 35 mm plastic dish with 500 µL of cell lysis buffer mixed with

protease and phosphatase inhibitors. Using a 1 mL micropipette, the FDB muscle bundle will be

triturated against the side of the dish to break up the fibers. Trituration will be repeated until the

muscle will be stripped from the surrounding white connective tissue. The FDB will be collected

in a 1.5 mL microcentrifuge tube and will be washed 3 times by gently resuspending muscle

fibers in 500 µL of cell lysis buffer with the protease cocktail. Each time, the tube will be

centrifuged at 3000 rpm and supernatant will be discarded. Muscle fibers will be homogenized in

300 µL of cell lysis buffer, protease cocktail added, and further processed to determine protein

concentration for western blotting analysis.

Blood sampling and flow cytometry analysis. To determine serum cytokine profile, blood

samples will be drawn using Microvette CB 300 Z tubes through the lateral saphenous vein

which runs dorsally and then laterally over the tarsal joint. Blood will be then centrifuged at

8000 rpm for 10 minutes at 4oC for serum separation. Serum samples will be blocked with mouse IL-6, IL-10, MCP-1, IFN-ϒ, and TNF-α capture beads using the commercial Cytometric

Bead Array (CBA) Mouse Inflammation Kit (cat# 552364) according to the manufacturer's instructions (BD Biosciences).

Wheel running protocol. mTLR4-/- and WT mice will undergo 4 weeks of voluntary running on

a cage-wheel protocol connected to the LabVIEW software, which monitors distance run,

running rate, and running duration. Wheel rotations with a diameter of 11.5 cm are counted

continuously, and the sum is sent to the software every two minutes.

Body composition analysis. Body mass will be determined prior to assessing body composition

using a portable balance (measuring to 0.01g). The body composition of mice will be assessed

using a Bruker LF90 NMR analyzer to measure lean and fat mass for all mice prior to and post

the wheel running program.

Testing aerobic capacity. Using incremental treadmill running, the testing protocol begins with

5 minutes warm-up at low work rate followed by stepwise increase in the power output. Mice

will go through incremental running starting at 0.20 m/s and 0˚ inclination. The treadmill

velocity will be increased by 0.03 m/s every two minutes until the mice reach exhaustion and

refuse to keep running. The speed at which the mouse cannot continue running will be recorded.

The testing session is to be terminated at any point at which a mouse meets either of these

exhaustion criteria:

1. Spending greater than 5 consecutive seconds on the shock grid without attempting to reengage

the treadmill.

2. Spending greater than 50% of its time on the shock grid.

3. The third time a mouse is willing to sustain 2 seconds or more of shock without attempting to

reengage the treadmill.

In-vivo plantar flexor torque assay. In-vivo measures of plantar flexor torque (PFT) will be obtained with mice under isoflurane anesthesia (platform at 35ºC). The mouse’s foot will be secured to a torque transducer (Aurora Scientific) and the tibial nerve will be stimulated at 2Hz for a series of contractions to fatigue (4 bouts X 8 minutes; 1 minute rest between bouts). At the end of the fatigue protocol, the leg will be aseptically cleaned with povidone-iodine solution while the mouse recovered from anesthesia (~2-5 minutes in 100% O2). Hindlimb PFT data will be analyzed using Dynamic Muscle Analysis (DMA) software versions 3.2 (Aurora Scientific).

Skeletal muscle whole homogenate preparation. Approximately 50 mg of fresh muscle

samples will be immediately placed into 0.2 ml of a modified sucrose EDTA medium (SET) on

ice containing 250 mM sucrose, 1 mM EDTA, 10 mM tris-HCl, and 1mM ATP, pH 7.4. Muscle

samples will be then minced with scissors and then SET buffer will be added to a 20-fold diluted

(wt:vol) suspension. The minced samples will be homogenized in a Potter-Elvehjem glass

homogenizer at 12 passes across 30 seconds at 150 rpm with a motor-driven teflon pestle, and

measures of fatty acid oxidation, glucose oxidation, pyruvate oxidation, and maximal enzyme

activities will be performed.

Substrate metabolism and metabolic flexibility. Red gastrocnemius and quadriceps muscles

will be collected for substrate metabolism. End products of substrate oxidation as measured by

[1-14C]-palmitic acid oxidation to 14CO2, [1-14C]-pyruvic acid oxidation to 14CO2 and [U-14C]-

glucose oxidation to 14CO2 will be assessed. Metabolic flexibility will be assessed by measuring

14 + [1- C]-pyruvate oxidation /- palmitic acid. For FAO, GO, and PDH activity assays, tissue

homogenates from red skeletal muscles will be incubated in 1μCi/mL of [1-14C]-palmitic acid,

1μCi/mL of [U-14C]-glucose, and 0.35 μCi/mL of [1-14C]-pyruvic acid, respectively, for 1 hour

and then acidified with 45% perchloric acid to elute gaseous CO2. 14CO2 will be trapped in 1M

sodium hydroxide for 1 hour. The sodium hydroxide will be then sampled and placed in a liquid

scintillation counter and quantified. Pyruvate oxidation will be used to assess the activity of

pyruvate dehydrogenase (PDH), the enzyme that catalyzes the oxidation of pyruvate resulting in

the provision of glucose-derived acetyl CoA to the TCA cycle. This is a direct measure of

pyruvate dehydrogenase activity as the number one carbon of pyruvate is liberated as CO2 in the

oxidation of pyruvate to acetyl-CoA.

Assessing glycolytic intermediates. For determination of muscle glucose and glucose 6-

phosphate concentrations, 300 μl of extracted muscle sample will be added to a tube containing

1.7 mL of 0.47 M triethanolamine, 35 mM MgCl2, 25 mM EDTA, and 1.2 mM NADP. An

aliquot (200 μl) will be then added in triplicate to a 96-well plate. Initial absorbance (OD1) will

be recorded using a spectrophotometer (SPECTRAmax ME, Molecular Devices Corporation,

Sunnyvale California) at a wavelength of 340 nm. Glucose-6-phosphate dehydrogenase (35 μl;

3.5 IU/sample) will be added to each tube and incubated for 20 min. Triplicate aliquots from each tube will be transferred to 96-well plates and absorbance will be measured again (OD2) at

340 nm. For determination of glucose, hexokinase (21 μl; 2.1 IU/sample) and ATP (41 μl; 11 mg/ml) will be added to each tube. Tubes will be allowed to incubate for 20 min before triplicate aliquots will be transferred to 96-well plates and absorbance (OD3) measured at 340 nm.

Absorbance values associated with muscle glucose 6-phosphate will be calculated by subtracting

OD1 from OD2. Similarly, absorbance values associated with muscle glucose will be determined by subtracting OD2 from OD3.

Phosphofructokinase (PFK) activity. PFK activity will be measured spectrophotometrically at

340nm at 37°C., Briefly, 30ul of sample homogenate will be pipette in duplicate (or triplicate)..

Then, assay buffer (12 mM MgCl2 , 400 mM KCL, 2 mM AMP, 1 mM ATP, 0.17 mM NADH,

0.0025 mg/mL, Antimycin 0.05 mg/mL Aldolase 0.05 mg/mL GAPDH, in100mM Tris buffer, pH=8.2) will be added into each well. After a 2 min background reading, 3 mM fructose-6-

phosphate will be added to each sample well and followed by a 7 minute kinetic reading.

Changes in absorbance across time will be recorded and expressed relative to protein content.

PFK activity will be calculated and reported as μmol/min/mg.

Citrate synthase (CS) activity. Citrate synthase catalyzes the formation of citrate and CoASH

from acetyl-CoA and oxaloactetate. CoASH reduces DTNB and CS activity will be determined from the reduction of DTMB over time. Briefly, ten microliters of a 1:5 diluted muscle homogenate will be added, in duplicate, to 170μl of a solution containing Tris buffer (0.1M, pH

8.3), DNTB (1mM, in 0.1M in Tris buffer) and oxaloacetate (0.01M, in 0.1M Tris buffer).

Following a 2min background reading, the spectrophotometer (SPECTRAmax ME, Molecular

Devices Corporation, Sunnyvale California) will be calibrated and 30μl of 3 mM acetyl CoA will be added to initiate the reaction. Absorbance will be measured at 405nm at 37C every 12 seconds for 7 minutes. CS activity will be calculated and reported as μmol/min/mg.

Malate dehydrogenase (MDH) activity. MDH activity will be measured spectrophotometrically at 340nm at 37°C. Briefly, 10ul of sample will be pipetted in duplicate or triplicate in wells. Then, 290ul of reaction media (0.1 M potassium phosphate buffer, PH 7.4 plus

0.006 M oxaloacetic acid, prepared in potassium phosphate buffer plus 0.00375 M NADH, prepared in potassium phosphate buffer) will be added to the wells and samples will be read for 5 minutes at 340nm. The rate of disappearance of NADH will be analyzed and expressed relative to protein content. MDH activity will be calculated and reported as μmol/min/mg.

β-hydroxyacyl-CoA dehydrogenase (β-HAD) activity. For the determination of β-HAD activity, oxidation of NADH to NAD will be measured. Volume of 35 μl of whole muscle homogenate will be added to 190μl of a buffer containing 0.1M liquid triethanolamine, 5mM

EDTA tetrasodium salt dihydrate, and 0.45mM NADH. The spectrophotometer (SPECTRAmax

PLUS 384, Molecular Devices Corporation, Sunnyvale California) will be calibrated and 15μl of

2mM acetoacetyl CoA will be added to initiate the reaction. Absorbance will be measured at 340

nm every 12 seconds for 5 minutes at 37C. β-HAD activity will be calculated and reported as

μmol/min/mg.

Cytochrome c oxidase (COX) activity. Cytochrome c oxidase enzyme activity will be measured spectrophotometrically at 550nm every 10 seconds for 7 minutes. Absorbance will be measured based on the oxidation of ferrocytochrome c to ferricytochrome c by cytochrome c oxidase. COX activity will be calculated and reported as μmol/min/mg.

RNA extraction and qRT-PCR. mRNA will be extracted using an RNeasy Mini Kit (Qiagen,

Valencia, CA) and DNase I treatment (Qiagen, Valencia, CA), according to the manufacturer's instructions. qRT-PCR will be performed using an ABI PRISM 7900 Sequence Detection

System instrument and TaqMan Universal PCR Master Mix will be used according to the manufacturer's specifications (Applied Biosystems, Foster City, CA). Target gene expression will be normalized to β-actin RNA levels. Primers and 5# FAM-labeled TaqMan probes will be purchased as prevalidated assays (Applied Biosystems, Foster City, CA).

Statistical analysis. Descriptive and inferential statistics methods will be employed to analyze

the collection of data of the study. The descriptive statistics as measured by mean and standard

error will be used to summarize the sample and to quantitatively describe the main features of the

data (e.g. body mass, lean mass, fat percent… etc.). Results are expressed as mean ± standard

error of the mean (SEM). As we are interested in studying the interaction effect of both exercise

and genotype differences (WT vs. mTLR4-/-) on manipulating exercise-induced myokine and metabolic responses, the statistical significances of differences among control and experimental groups will be determined using the Two-Way ANOVA. If a significant F-ratio will be obtained, comparisons between groups will be made using Tukey’s post-hoc test. Furthermore, Student’s t-

test will be used to report the differences in gain scores between the control and experimental groups to better understand the genotype X exercise interaction effect. The level of significance will be established as P < 0.05.

CHAPTER V

Muscle-Specific Deletion of Toll-like Receptor-4 Impairs Metabolic Adaptation to Wheel Running in Mice.

Abstract

Toll like receptor 4 (TLR4) is a transmembrane inflammatory receptor expressed ubiquitously on

the cell surface of skeletal muscle and other metabolic tissues. Skeletal muscle gains favorable

inflammatory-mediated metabolic adaptations from exercise training. However, overexpression

as well as pathogen-stimulated activation of muscle TLR4 (mTLR4) and subsequent downstream

inflammatory signaling can compromise skeletal muscle metabolism. This paradox raises a

concern about the significance of mTLR4 signaling to the metabolic gains associated with

exercise performance. Thus, herein, we aimed to investigate the role of mTLR4 in modulating

exercise-induced metabolic adaptations. To this end, we developed a novel muscle-specific

TLR4 knockout (mTLR4-/-) mouse model on C57BL/6JJ background. Male mTLR4-/- mice and

wild type (WT) littermates maintained either regular cage activity or performed voluntary wheel

running for 4 weeks. Following the training protocol, gastrocnemius and quadriceps muscles

were harvested. Notably, mTLR4 deletion revealed marked reductions in downstream

interleukin-1 receptor-associated kinase-4 (IRAK4) phosphorylation in muscle homogenates of

mTLR4-/- mice compared to WT littermates. In addition, disruption of mTLR4 signaling

completely blunted the metabolic adaptations observed in trained WT mice. In particular, trained

WT mice significantly increased levels of glucose oxidation (GO) and fatty acid oxidation

(FAO) coupled with a substantial increase in glycolytic substrate availability as well as

improvements in pyruvate dehydrogenase (PDH) activity, β-hydroxyacyl-CoA dehydrogenase

(β-HAD), and citrate synthase (CS) catalytic capacities. In contrast, no significant differences

were detected in levels of substrate metabolism or enzymatic activity in trained mTLR4-/- mice

compared with their sedentary controls. Taken together, this study introduces a novel muscle-

specific TLR4 knockout mouse model. Furthermore, our findings identify mTLR4 as a major

immunomodulatory effector of exercise-induced metabolic adaptations in skeletal muscle.

Key Words: skeletal muscle, toll-like receptor-4, wheel running, glucose oxidation, fatty acid

oxidation.

Introduction

Skeletal muscle is a major metabolic tissue that accounts for approximately 40-50 % of total

body mass [1]. Toll-like receptor 4 (TLR4) is an inflammatory signaling receptor that is widely

expressed in skeletal muscle and other metabolic tissues [2, 3]. TLR4 is not only known to

stimulate the production of inflammatory cytokines, but also modulates carbohydrate and lipid

metabolism [4, 5]. Many reports have indicated that skeletal muscle is the source of elevations

observed in plasma cytokines during and after exercise [6, 7]. Muscle-derived cytokines

(myokines) released in response to exercise such as interleukin-6 (IL-6), monocyte

chemoattractant protein (MCP-1), interleukin 8 (IL-8), and interleukin 15 (IL-15), exhibit varied

time course profiles [8, 9], suggesting involvement of different regulatory mechanisms. Several

of these myokines that are produced with muscle contraction, most notably IL-6, mediate favorable metabolic responses to exercise [10, 11]. In vitro exposure to IL-6 has been shown to

stimulate glycogen synthesis and glucose oxidation in human skeletal muscle [12]. It has also

been observed that IL-6 infusion in humans augments lipid oxidation with increased reliance on

free fatty acids as a source for energy production in skeletal muscle under basal conditions [13].

Of note, many of these exercise-responsive inflammatory mediators of metabolic events appear to be direct targets for the upstream TLR4 signals. However, to the best of our knowledge, it has 41

not been determined whether mTLR4 specific intracellular signaling transduction is necessarily

involved in regulating the metabolic responses to exercise. Thus the role of TLR4 as a potential

modulator of exercise-induced metabolic adaptations is appropriate to study.

Recent research has documented that prolonged exercise stimulates the induction of molecules

including heat shock proteins (HSPs) and plasma free fatty acids (FFAs) that are suggested to be

activators of certain toll-like receptors (TLRs) and the subsequent downstream cytokine

production [14, 15]. Additionally, similar to an endotoxin challenge (e.g. lipopolysaccharide,

LPS), a down-regulation in TLR4 gene expression has been observed following prolonged vigorous exercise [16, 17]. These intriguing results have developed a recent interest postulating a role for TLRs in modulating inflammatory events and presumably metabolic adaptations in response to exercise [18]. In line with this notion, non-esterified fatty acids (NEFA) have been suggested to activate P38 MAPK and JNK pathways via TLR4-dependent mechanisms in skeletal muscle during endurance exercise [19]. To date, however, there is no direct evidence to support this paradigm. Furthermore, the vast majority, if not all, of related studies emphasize the suppressive actions of exercise training on TLR4 abundance and activation. We adopted a novel concept hypothesizing that mTLR4 signaling is an essential contributor to the metabolic gains of long-term exercise. Therefore the objective of the present study was to investigate the role of mTLR4 signaling in modulating skeletal muscle metabolic response to exercise training.

Material and Methods

Animal husbandry and generation of mTLR4-/- mice. Animal studies were performed under

an approved protocol by the Institutional Animal Care and Use Committee (IACUC) at Virginia

Tech. The muscle-specific TLR4 knockout (mTLR4-/-) mouse model on C57BL/6J background

was created at the Transgenic Core at the Pennington Biomedical Research Center using the Cre- 42

mediated recombination driven by the muscle creatine kinase promoter. Male mTLR4-/- and wild type (WT) littermates were maintained on a 12-hour light/dark cycle and fed a standard chow diet. All experiments were performed following an overnight fast (10-12 hours), unless otherwise stated. To confirm muscle-specific deletion of TLR4, animals were sacrificed and flexor digitorum brevis (FDB) muscle was isolated. The FDB muscle group was used because isolation of single muscle fibers limits confounding influences of non-muscle cell types that are present in whole muscle groups (e.g. gastrocnemius and quadriceps).

FDB muscle fibers isolation and preparation. Mice were euthanized using CO2 asphyxiation followed by cervical dislocation according to Virginia Tech IACUC protocols. The hind foot was cut off above the ankle joint. The foot was pinned with the plantar side facing up onto a flat foam board. Skin was reflected starting from the proximal side of the foot, at the ankle, and was pulled upwards toward the toes. The FDB muscle bundle was dissected by cutting the tendon at the heel and pulling it upward while detaching the muscle fibers from other adjacent tissues. The

FDB muscle bundle was removed by cutting at the insertion tendons attached to each of the four lateral toes. The FDB muscle was immediately placed in a tube with 500 µL of dissociation buffer containing cell lysis buffer mixed with protease and phosphatase inhibitors (1/100) as well as collagenase (4 mg/mL). The tube was rocked in a horizontal position on a shaker for approximately 90 minutes at 37o C. Digested muscle was transferred into a 35 mm plastic dish

with 500 µL of cell lysis buffer mixed with protease and phosphatase inhibitors. Using a 1 mL

micropipette, the FDB muscle bundle was triturated against the side of the dish to break up the fibers. Trituration was repeated until the muscle was stripped from the surrounding white connective tissue. The FDB was collected in a 1.5 mL microcentrifuge tube and washed 3 times

by gently resuspending muscle fibers in 500 µL of cell lysis buffer with the protease cocktail.

Each time, the tube was centrifuged at 3000 rpm and supernatant was discarded. Muscle fibers were homogenized in 300 µL of cell lysis buffer, protease cocktail added, and further processed to determine protein concentration for western blotting analysis.

Western blot analysis. Western blot analysis was performed using FDB muscle fibers. Samples were homogenized in 50 mM Hepes, (pH7.5), 15 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 2 mM

EDTA, 10% glycerol, 1 % Triton x-100, 10 mM NaP2O7, 100 mM NaFl, 10 mM PMSF, and 10 g/mL aprotinin. Proteins (~30ug per sample) were separated using a 10% Criterion-Tris-HCl gel

(Bio-Rad, Hercules, CA) and subsequently transferred to PVDF membrane (Bio-Rad, Hercules,

CA). PVDF membrane were probed with a primary antibody against TLR4 (abcam, cat# ab13556) in a 1:200 dilution of BSA, phospho-IRAK4 (Cell Signaling, cat# 7652S) in a 1:1000 dilution of BSA.

Wheel running protocol. mTLR4-/- and WT mice underwent 4 weeks of voluntary running on a cage-wheel protocol connected to the LabVIEW software, which monitors distance run, running rate, and running duration. Wheel rotations with a diameter of 11.5 cm were counted continuously, and the rotations were updated by the software every two minutes.

Body composition analysis. Body mass was determined prior to assessing body composition using a portable balance (measuring to 0.01g). We assessed the body composition of mice using a Bruker LF90 NMR analyzer to assess lean and fat mass for all mice prior to and following the wheel running program.

Skeletal muscle whole homogenate preparation for substrate metabolism. Approximately 50 mg of fresh muscle samples were immediately placed into 0.2 ml of a modified sucrose EDTA medium (SET) on ice containing 250 mM sucrose, 1 mM EDTA, 10 mM tris-HCl, and 1mM

ATP, pH 7.4. Muscle samples were then minced with scissors and SET buffer was added to

achieve a 20-fold diluted (wt:vol) suspension. The minced samples were homogenized in a

Potter-Elvehjem glass homogenizer at 12 passes across 30 seconds at 150 rpm with a motor-

driven teflon pestle, and measures of fatty acid oxidation, glucose oxidation, pyruvate oxidation,

and maximal enzyme activities were performed.

Substrate metabolism and metabolic flexibility. Red gastrocnemius and quadriceps muscles

were collected for substrate metabolism. End products of substrate oxidation as measured by [1-

14C]-palmitic acid oxidation to 14CO2, [1-14C]-pyruvic acid oxidation to 14CO2 and [U-14C]- glucose oxidation to 14CO2 were assessed. Metabolic flexibility was assessed by measuring [1-

14 + C]-pyruvate oxidation /- 100uM palmitic acid. For FAO, GO, and PDH activity assays, tissue

homogenates from red skeletal muscles were incubated in 1μCi/mL of [1-14C]-palmitic acid,

1μCi/mL of [U-14CO2]-glucose, and 0.35 μCi/mL of [1-14C]-pyruvic acid, respectively, for 1

hour and then acidified with 45% perchloric acid to elute gaseous CO2. 14CO2 was trapped in

1M sodium hydroxide for 1 hour. The sodium hydroxide was then sampled and placed in a liquid

scintillation counter and quantified. Pyruvate oxidation was used to assess the activity of

pyruvate dehydrogenase (PDH), the enzyme that catalyzes the oxidation of pyruvate resulting in

the provision of glucose-derived acetyl CoA to the TCA cycle. This is a direct measure of

pyruvate dehydrogenase activity as the number one carbon of pyruvate is liberated as CO2 in the

oxidation of pyruvate to acetyl-CoA.

Assessing glycolytic intermediates. For determination of muscle glucose and glucose 6-

phosphate concentrations, 300 μl of extracted muscle sample were added to a tube containing 1.7

mL of 0.47 M triethanolamine, 35 mM MgCl2, 25 mM EDTA, and 1.2 mM NADP. An aliquot

(200 μl) was then added in triplicate to a 96-well plate. Initial absorbance (OD1) was recorded

using a spectrophotometer (SPECTRAmax ME, Molecular Devices Corporation, Sunnyvale

California) at a wavelength of 340 nm. Glucose-6-phosphate dehydrogenase (35 μl; 3.5

IU/sample) was added to each tube and incubated for 20 min. Triplicate aliquots from each tube were transferred to 96-well plates and absorbance was measured again (OD2) at 340 nm. For determination of glucose, hexokinase (21 μl; 2.1 IU/sample) and ATP (41 μl; 11 mg/ml) were added to each tube. Tubes were allowed to incubate for 20 min before triplicate aliquots were transferred to 96-well plates and absorbance (OD3) measured at 340 nm. Absorbance values associated with muscle glucose 6-phosphate were calculated by subtracting OD1 from OD2.

Similarly, absorbance values associated with muscle glucose were determined by subtracting

OD2 from OD3.

Citrate synthase (CS) activity. Citrate synthase catalyzes the formation of citrate and CoASH from acetyl-CoA and oxaloactetate. CoASH reduces DTNB and CS activity was determined from the reduction of DTMB over time. Briefly, ten microliters of a 1:10 diluted muscle homogenate was added, in duplicate, to 170μl of a solution containing Tris buffer (0.1M, pH

8.3), DNTB (1mM, in 0.1M in Tris buffer) and oxaloacetate (0.01M, in 0.1M Tris buffer).

Following a 2min background reading, the spectrophotometer (SPECTRAmax ME, Molecular

Devices Corporation, Sunnyvale California) was calibrated and 30μl of 3 mM acetyl CoA was added to initiate the reaction. Absorbance was measured at 405nm at 37C every 12 seconds for 7 minutes. CS activity was calculated and reported as μmol/min/mg.

β-hydroxyacyl-CoA dehydrogenase (β-HAD) activity. For the determination of β-HAD activity, oxidation of NADH to NAD was measured. 35 μl of whole muscle homogenate was added to 190μl of a buffer containing 0.1M liquid triethanolamine, 5mM EDTA tetrasodium salt dihydrate, and 0.45mM NADH. The spectrophotometer (SPECTRAmax PLUS 384, Molecular

Devices Corporation, Sunnyvale California) was calibrated and 15μl of 2mM acetoacetyl CoA was added to initiate the reaction. Absorbance was measured at 340 nm every 12 seconds for 5 minutes at 37C. β-HAD activity was calculated and reported as μmol/min/mg.

Phosphofructokinase (PFK) activity. PFK activity was measured spectrophotometrically at

340nm at 37°C., Briefly, 30ul of sample homogenate were pipette in duplicate (or triplicate)..

Then, assay buffer (12 mM MgCl2 , 400 mM KCL, 2 mM AMP, 1 mM ATP, 0.17 mM NADH,

0.0025 mg/mL, Antimycin 0.05 mg/mL Aldolase 0.05 mg/mL GAPDH, in100mM Tris buffer, pH=8.2) was added into each well. After a 2 min background reading, 3 mM fructose-6- phosphate was added to each sample well and followed by a 7 minute kinetic reading. Changes in absorbance across time were recorded and expressed relative to protein content. PFK activity was calculated and reported as μmol/min/mg.

RNA extraction and qRT-PCR. mRNA was extracted using an RNeasy Mini Kit (Qiagen,

Valencia, CA) and DNase I treatment (Qiagen, Valencia, CA), according to the manufacturer's instructions. qRT-PCR was performed using an ABI PRISM 7900 Sequence Detection System instrument and TaqMan Universal PCR Master Mix used according to the manufacturer's specifications (Applied Biosystems, Foster City, CA). Target gene expression was normalized to

β-actin RNA levels. Primers and 5# FAM-labeled TaqMan probes were purchased as prevalidated assays (Applied Biosystems, Foster City, CA).

Statistical analysis. Descriptive and inferential statistics methods were employed to analyze the collection of data of the study. Descriptive statistics as measured by mean and standard error were used to summarize the sample and to quantitatively describe the main features of the data

(e.g. body mass, lean mass, fat percent… etc.). Results are expressed as mean ± standard error of

the mean (SEM). As we were interested in studying the interaction effect of both exercise and

genotype differences (WT vs. mTLR4-/-) on manipulating exercise-induced myokine and metabolic responses, the statistical significances of differences among control and experimental groups were determined using the Two-Way ANOVA. If a significant F-ratio was obtained, comparisons between groups were made using Tukey’s post-hoc test. Furthermore, Student’s t- test was used to report the differences in gain scores between the control and experimental groups to better understand the genotype X exercise interaction effect. The level of significance was established as P < 0.05.

Results mTLR4-/- mouse model. mTLR4-/-mice and their WT littermates were identified following tail

biopsy genotyping procedures. mTLR4-/- mice showed 68% lower FDB TLR4 protein content

compared to the WT counterparts (Figure s 1A, B). Investigating key downstream markers

mediating the TLR4 signaling transduction, mTLR4-/- mice exhibited ~ 70% lower

phosphorylation of IRAK4 (P < 0.05) (Figures 1C, D).

Figure 1. mTLR4 protein content is decreased and is coupled with lower IRAK4

phosphorylation in mTLR4-/- mice. WT (n=6) and mTLR4-/- mice (n=6) were sacrificed after

overnight fasting and muscle lysates were processed for western blotting analysis to assess

muscle TLR4 protein abundance (A, B) as well as IRAK4 phosphorylation status (C, D),

downstream from TLR4. Data are expressed as mean ± SEM (*P < 0.05).

Voluntary wheel running and body composition parameters. Upon the completion of the wheel running program, no differences were detected in voluntary wheel running performance

(WT: 1117 ± 246 meter/day vs. mTLR4-/-: 1357 ± 417 meter/day) between WT and mTLR4-/-

mice (Figure 2A). Meanwhile, wheel running was associated with significant reductions in body

mass and fat mass as well as an increase in fat free mass in both WT and mTLR4-/- mice (P <

0.05) (Figures 2B-D).

Figure 2. Wheel running was associated with reductions in body mass and fat mass as well as an increase in fat free mass, however, no genotype differences were observed in wheel running performance. WT (n=6) and mTLR4-/- mice (n=7) underwent 4 weeks of voluntary

running on a cage-wheel protocol connected to LabVIEW software, which monitored distance

run, running rate, and running duration. Average daily wheel running performance was

determined (A). Prior to the wheel running protocol, body composition parameters (B-D) were assessed via the Bruker mini spec LF90. Independent from genotype, wheel running reduced

body mass (B), fat mass (C), and fat free mass (D). Data are presented as means ± SEM (*P <

0.05).

mTLR4 and wheel running-induced metabolic adaptations. Data from whole red skeletal

muscle homogenates indicate genotype-specific metabolic improvements in response to wheel running. Although in sedentary conditions the basal metabolic substrate oxidation was not different between WT and mTLR4-/- mice, results showed a significant genotype-wheel running interaction as manifested by a greater change in oxidative capacity of WT mice compared with mTLR4-/- littermates. Compared to respective sedentary controls, wheel running resulted in

28.3% higher fatty acid oxidation in WT mice (WT Sed: 68.4 ± 8.1 nmol/mg protein/hr vs. WT

WR: 87.7 ± 5.5 nmol/mg protein/hr) and 16 % lower oxidation in mTLR4-/- mice (mTLR4-/- Sed:

75 ± 4.8 nmol/mg protein/hr vs. mTLR4-/- WR: 63 ± 3.4 nmol/mg protein/hr) (Figures 3A, B;

P<0.05). Moreover, compared to sedentary controls, wheel running induced 50.7 % higher

glucose oxidation in WT mice (WT Sed: 33 ± 2.9 nmol/mg protein/hr vs. WT WR: 49.7 ± 3.1

nmol/mg protein/hr: P<0.05) and 15.7% lower oxidation in mTLR4-/- mice (mTLR4-/- Sed: 35.5

± 2.8 nmol/mg protein/hr vs. mTLR4-/- WR: 30.4 ± 3.2 nmol/mg protein/hr) (Figures 3C, D;

P<0.05). Despite the substantial differential metabolic response to wheel running between WT

and mTLR4-/- mice, differences in metabolic flexibility did not reach significant levels in trained

WT and mTLR4-/- mice compared to their respective sedentary controls (P = 0.09, Figures 3E,

F).

Figure 3. mTLR4 mediates wheel running-induced increases in substrate oxidation in skeletal muscle. WT (n=12) and mTLR4-/- mice (n=14) were sacrificed 24 hours after the

completion of the wheel running protocol and red gastrocnemius and quadriceps muscle were

collected for substrate metabolism. Fatty acid oxidation (A, B), glucose oxidation (C, D), and metabolic flexibility (E, F) were assessed. To assess glucose oxidation, muscle homogenates were incubated in [U-14C]-glucose at concentrations of 1µCi/mL and 14CO2 production was

measured. For fatty acid oxidation, muscle homogenates were incubated in 1µCi/mL of [1-14C]- palmitic acid and 14CO2 production was measured. Data are presented as means ± SEM (*P <

0.05). mTLR4 and wheel running-induced adaptations of muscle enzymatic machinery. Increased oxidative metabolism observed in trained WT mice was supported by functional augmentations in enzymatic activity of PDH (WT Sed: 1880.4 ± 121.5 nmol/mg protein/hr vs. WT WR: 2658.1

± 210.5 nmol/mg protein/hr: P<0.05) (Figure 4A, B), CS (WT Sed: 409.5 ± 24 µm/mg/min vs.

WT WR: 578.6 ± 28.5 µm/mg/min P<0.05) (Figure 4C, D) and β-HAD (WT Sed: 162.4 ± 9.6

µm/mg/min vs. WT WR: 213 ± 16.3 µm/mg/min: P<0.05) (Figure 4E, F). Although PFK enzymatic activity trended to an increase in WT mice and to a decrease in the mTLR4-/- littermates, these differences did not achieve statistical significance (Figure 4G, H). Meanwhile, muscle glucose content remained unchanged between sedentary control and trained groups in

WT and mTLR4-/- mice (Figure 5A). However, wheel running induced a significant increase in muscle glucose-6-phosphate content in trained WT mice (WT Sed: 36.4 ± 5.3 µmol/mg vs. WT

WR: 83.2 ± 5.5 µmol/mg: P<0.05) (Figure 5B). On the other hand, the mTLR4-/- littermates failed to develop any of the aforementioned oxidative adaptations displayed by WT mice in response to wheel running.

Figure 4. mTLR4 modulates the oxidative capacity of muscle mitochondrial enzymes in response to wheel running. WT (n=12) and mTLR4-/- mice (n=14) were fasted overnight and

whole muscle homogenates were obtained to measure mitochondrial enzymatic activity.

Pyruvate dehydrogenase activity (A, B) was measured by incubating muscle homogenates in [1-

14C]-Pyruvic acid at concentrations of 0.35μCi/mL and 14CO2 production was measured. Citrate

synthase (C, D), β-hydroxyacyl-CoA dehydrogenase (E, F), and phosphofruvtokinase-1 (G, H)

activities were determined spectrophotometrically. Wheel running-induced oxidative adaptations

were completely blunted in mTLR4-/- mice compared to WT littermates. Data are presented as

means ± SEM (*P < 0.05).

Figure 5. mTLR4 signaling is required to enrich glucose-6-phosphate metabolite pools in

response to wheel running. The glycolytic metabolites, glucose and glucose-6-phosphate, were

measured spectrophotometrically in muscle extracts from WT (n=12) and mTLR4-/- mice (n=14).

Across different genotypes, wheel running did not impact intracellular glucose levels (A).

However, G-6-P availability was significantly enhanced in trained WT mice but not in mTLR4-/-

littermates (B). Data are presented as means ± SEM (*P < 0.05).

Gene expression in response to wheel running. Wheel running did not induce significant

changes in mRNA levels of PGC1-α (Figure 6A), PPAR-delta (Figure 6B), and IL-6 (Figure 6C) in WT or mTLR4-/- mice.

Figure 6. Changes in gene expression of key metabolic and inflammatory regulators in response to wheel running. Across different genotypes, PGC1-α (A), PPAR-delta (B), and IL-6

(C) mRNA levels remained unchanged after wheel running, when comparing the sedentary WT and mTLR4-/- mice to their respective trained groups. Data were normalized to β-Actin and presented as means ± SEM.

Discussion

Physical exercise induces multi-systemic changes that excite global mobilization of signaling

molecules important for cell communication and survival. These signaling messengers are

detected by target cells via membrane receptors translating external information into intercellular

responses. In turn, host cells adjust to new demands imposed by this exercise stimulus. TLR4 has

consistently been portrayed as a pleiotropic cellular receptor modulating multiple inflammatory

and metabolic pathways [20-22]. Nonetheless, the significance of TLR4 in exercise metabolism

has not been revealed. Identifying the role of TLR4 in exercise performance and energy control

may delineate a mechanistic basis for the disrupted exercise-induced adaptations often observed

in inflammatory conditions [23-25] characterized with pathologically altered TLR4 signaling,

such as obesity and type 2 diabetes [26]. In this study, we used a wheel running exercise

modality to investigate the role of skeletal muscle TLR4 in voluntary exercise performance and

the subsequent metabolic adaptations. We utilized mTLR4-/- mice and wild type littermates

which underwent 4 weeks of voluntary wheel running. Following the completion of the exercise

protocol, skeletal muscle was collected for oxidative metabolism measurements and assessment of enzymatic activity and gene expression of protein targets thought to be important regulators of metabolic pathways. Herein, we provide data that suggest a possible role for mTLR4 in modulating molecular events required to initiate and develop a normal metabolic adaptation in oxidative tissues in response to exercise training.

For almost two decades since the discovery of TLR4, there has been a growing interest in

exploring its implications in health and disease. The loss of function approach has provided a

great tool for examining the physiological role of TLR4 within different populations and

conditions. Many laboratories have developed various TLR4 knockout mouse models for their

own research [27-30]. As many of the mouse models are whole body knockouts of TLR4

signaling, it is unclear which exact cell type(s) mediate the phenotypes observed in these models.

Distinguishable from others, we selectively generated a targeted knockout for mTLR4 in the

genome of C57BL/6J mice. Our model manifests a 70% reduction in mTLR4 protein levels. The

marginal TLR4 residual protein detected in muscle homogenates from mTLR4-/- mice could plausibly be attributed to other cell types inherent to muscle microenvironment such as macrophages and dendritic cells [31, 32]. To further confirm the functional inactivation of mTLR4, we subsequently analyzed the phosphorylation status of IRAK4 which has been identified as a downstream obligatory contributor to TLR-dependent immune responses [33].

Although IRAK4 activation is a target event of other upstream membrane-spanning receptors

[34], IRAK4 protein phosphorylation was decreased by roughly 60% in mTLR4-/- mice

compared with WT littermates. As expected, TLR4 protein content and the downstream signal

transductions were specifically disrupted in mTLR4-/- mice. However, the placement of our gene mutagen construct under the control of the creatine kinase promoter could raise a tissue- specificity concern. The disruption of TLR4 signaling in cardiac muscle is a potential limitation to heart functionality and therefor to the exercise capacity of mTLR4-/- mice. Noteworthy,

previous studies have reported that C57BL/6J mice, out of 7 different inbred mouse strains, were

the highest performers on wheel running [35]. We speculated that voluntary, low intensity, wheel

running does not necessarily constitute a physical challenge to mTLR4-/- mice on the C57BL/6J

background. In concordance with this assumption, our WT and mTLR4-/- littermates exhibited similar exercise patterns with no significant differences in wheel running performance.

Wheel running has repeatedly been demonstrated to improve the oxidative capacity in rodents

[36-38]. Interestingly, typical exercise-induced metabolic adaptations were blocked as a result of

mTLR4 absence. Contrary to WT littermates, mTLR4-/- mice failed to enhance levels of fatty

acid oxidation after four weeks of wheel running training. These metabolic ramifications of

mTLR4 deficiency were further confirmed by the lack of wheel running-induced augmentations

of beta oxidation as measured by β-HAD enzymatic activity thereby limiting substrate flux

feeding into the tricarboxylic acid (TCA) cycle. TLR4 has previously been linked to key

regulators of cellular energy homeostasis. Our results are consistent with a recent report demonstrating that exercise-induced p38 MAPK signaling and JNK activation, two crucial modulators of fat metabolism, were abolished in TLR4-deficient mice [19]. Additionally, TLR4 deletion has been shown to diminish exercise-associated inflammatory response in a tissue-

specific manner, which implies impaired cytokine signaling [39]. Upon muscle contraction, IL6

release has been well documented to promote AMPK activation and the concomitant

accelerations in rates of fatty acid oxidation [11, 40]. These oxidative adaptations are possibly

depressed in mTLR4-/- mice. This paradigm represents a legitimate molecular basis for the role

of mTLR4 in modulating fat metabolism in response to exercise training. Furthermore, our data

indicate that carbohydrate utilization in response to exercise is also modulated by mTLR4

signaling. Improvements in glucose oxidation observed in WT mice were completely blunted by

the mTLR4 gene disruption. Undoubtedly, TLRs, including TLR4, up-regulate glycolysis in an

almost Warburg Effect fashion [41]. A primary target for TLR4-mediated elevation of glycolysis

is hexokinase [42]. Consequently, inhibiting mTLR4 is very likely inhibiting its activation of

glycolysis. Besides, as mentioned previously, TLR4-stimulated cytokine production includes IL-

6 which has recently been shown to enhance glycolysis through hexokinase- and

phosphofructokinase-dependent mechanisms [43]. In concert with these notions, levels of

upstream glycolytic intermediates remained unchanged in mTLR4-/- muscle, whereas WT

littermates exhibited a substantial increase in levels of glucose-6-phosphate in response to wheel running. Increased glucose uptake and glycogen reserves potentially underlie elevations observed in substrate availability in trained WT mice as oppose to mTLR4-/- counterparts.

Intriguingly, although not statistically significant, wheel running tended to increase

phosphofructokinase activity in WT mice, yet, trained mTLR4-/- littermates barely sustained

basal activity levels detected in their sedentary controls. It is important to bear in mind that

amplified beta-oxidation observed in trained WT mice may expectedly increase cytosolic citrate

content which could partially account for the restrained phosphofructokinase activity. These

findings offer a possible mechanistic understanding to our observation of a significant suppression of PDH adaptive response in mTLR4-/- mice, which failed to match PDH elevations

witnessed in WT mice in response to wheel running. Indeed, impaired glycolytic flux observed

in mTLR4-/- mice may limit substrate availability for mitochondria, thus abrogating the need for

PDH complex up-regulation. Taken together, concurrent maladaptation of key enzymes involved

in both beta-oxidation and aerobic glycolysis suggests a scarcity of acetyl CoA supply. Lack of

carbon substrates feeding into the TCA cycle may account for exercise-irresponsiveness of CS

activity elicited in trained mTLR4-/- mice compared with the corresponding WT littermates.

Noteworthy, however, the metabolic superiority of WT mice to mTLR4-/- littermates, as defined

by enzymatic capacity and substrate oxidation, was insufficient to induce noticeable phenotypic

differences in wheel running performance. Several factors can modulate maximum oxygen

consumption and increased aerobic capacity associated with exercise performance. Yet, the

contribution of mitochondrial enzymes to exercise-induced VO2 max adjustments is

controversial. Saltin and colleagues indicated that skeletal muscle has a propensity to overbuild

its enzymatic machinery to an extent that exceeds the oxygen transport capacity of the circulation

[44]. As such, the genotype discrepancies we found in enzyme capacities may not be a vital

determinant controlling submaximal exercise performance.

In conclusion, this study introduces a novel muscle-specific TLR4 knockout mouse model. In

addition, the current results reveal a vital role for mTLR4 in mediating exercise-induced

metabolic adaptations. Nevertheless, the absence of adaptations as observed in WT mice did not

impede voluntary running wheel performance in the mTLR4-/- mice. However, the underling

mechanisms linking exercise performance to mTLR4 activation and the downstream signaling

are yet to be fully elucidated. Moreover, controlled studies with known exercise doses are needed

for more reliable genotype comparisons.

References

1. Nader, G.A., Molecular determinants of skeletal muscle mass: getting the "AKT" together. Int J Biochem Cell Biol, 2005. 37(10): p. 1985-96. 2. Frost, R.A., G.J. Nystrom, and C.H. Lang, Lipopolysaccharide regulates proinflammatory cytokine expression in mouse myoblasts and skeletal muscle. Am J Physiol Regul Integr Comp Physiol, 2002. 283(3): p. R698-709. 3. Song, M.J., et al., Activation of Toll-like receptor 4 is associated with insulin resistance in adipocytes. Biochem Biophys Res Commun, 2006. 346(3): p. 739-45. 4. McMillan, R.P., et al., Selective overexpression of Toll-like receptor-4 in skeletal muscle impairs metabolic adaptation to high-fat feeding. Am J Physiol Regul Integr Comp Physiol, 2015. 309(3): p. R304-13. 5. Frisard, M.I., et al., Toll-like receptor 4 modulates skeletal muscle substrate metabolism. American Journal of Physiology-Endocrinology and Metabolism, 2010. 298(5): p. E988- E998. 6. Pedersen, B.K., Muscles and their myokines. The Journal of experimental biology, 2011. 214(2): p. 337-346. 7. Febbraio, M.A. and B.K. Pedersen, Muscle-derived interleukin-6: mechanisms for activation and possible biological roles. The FASEB Journal, 2002. 16(11): p. 1335- 1347. 8. Louis, E., et al., Time course of proteolytic, cytokine, and myostatin gene expression after acute exercise in human skeletal muscle. Journal of applied physiology, 2007. 103(5): p. 1744-1751.

9. Schild, M., et al., Effects of Acute Endurance Exercise on Plasma Protein Profiles of Endurance-Trained and Untrained Individuals over Time. Mediators of Inflammation, 2016. 2016: p. 11. 10. Smith, L.L., Cytokine hypothesis of overtraining: a physiological adaptation to excessive stress? Med Sci Sports Exerc, 2000. 32(2): p. 317-31. 11. Kelly, M., et al., AMPK activity is diminished in tissues of IL-6 knockout mice: the effect of exercise. Biochem Biophys Res Commun, 2004. 320(2): p. 449-54. 12. Glund, S., et al., Interleukin-6 directly increases glucose metabolism in resting human skeletal muscle. Diabetes, 2007. 56(6): p. 1630-1637. 13. Wolsk, E., et al., IL-6 selectively stimulates fat metabolism in human skeletal muscle. American Journal of Physiology-Endocrinology and Metabolism, 2010. 299(5): p. E832- E840. 14. Kaisho, T. and S. Akira, Toll-like receptor function and signaling. Journal of Allergy and Clinical Immunology, 2006. 117(5): p. 979-987. 15. Nguyen, M.A., et al., A subpopulation of macrophages infiltrates hypertrophic adipose tissue and is activated by free fatty acids via Toll-like receptors 2 and 4 and JNK- dependent pathways. Journal of Biological Chemistry, 2007. 282(48): p. 35279-35292. 16. Oliveira, A.G., et al., Physical exercise reduces circulating lipopolysaccharide and TLR4 activation and improves insulin signaling in tissues of DIO rats. Diabetes, 2011. 60(3): p. 784-96. 17. Fernandez-Gonzalo, R., et al., TLR4-mediated blunting of inflammatory responses to eccentric exercise in young women. Mediators Inflamm, 2014. 2014: p. 479395. 18. Gleeson, M., et al., The anti-inflammatory effects of exercise: mechanisms and implications for the prevention and treatment of disease. Nature Reviews Immunology, 2011. 11(9): p. 607-615. 19. Zbinden-Foncea, H., et al., TLR2 and TLR4 activate p38 MAPK and JNK during endurance exercise in skeletal muscle. Med Sci Sports Exerc, 2012. 44(8): p. 1463-72. 20. Pang, S., et al., Regulation of fasting fuel metabolism by toll-like receptor 4. Diabetes, 2010. 59(12): p. 3041-8. 21. Frisard, M.I., et al., Toll-like receptor 4 modulates skeletal muscle substrate metabolism. Am J Physiol Endocrinol Metab, 2010. 298(5): p. E988-98. 22. Takeda, K., T. Kaisho, and S. Akira, Toll-like receptors. Annu Rev Immunol, 2003. 21: p. 335-76. 23. Scheen, A.J., et al., Metabolic adaptation to prolonged exercise in severely obese subjects. Int J Obes, 1983. 7(3): p. 221-9. 24. Perez-Martin, A., et al., Balance of substrate oxidation during submaximal exercise in lean and obese people. Diabetes Metab, 2001. 27(4 Pt 1): p. 466-74. 25. Thyfault, J.P., et al., Impaired plasma fatty acid oxidation in extremely obese women. Am J Physiol Endocrinol Metab, 2004. 287(6): p. E1076-81. 26. Reyna, S.M., et al., Elevated toll-like receptor 4 expression and signaling in muscle from insulin-resistant subjects. Diabetes, 2008. 57(10): p. 2595-602. 27. Shi, H., et al., TLR4 links innate immunity and fatty acid-induced insulin resistance. J Clin Invest, 2006. 116(11): p. 3015-25. 28. Kim, F., et al., Toll-like receptor-4 mediates vascular inflammation and insulin resistance in diet-induced obesity. Circ Res, 2007. 100(11): p. 1589-96.

29. Saberi, M., et al., Hematopoietic cell-specific deletion of toll-like receptor 4 ameliorates hepatic and adipose tissue insulin resistance in high-fat-fed mice. Cell Metab, 2009. 10(5): p. 419-29. 30. Davis, J.E., et al., Tlr-4 deficiency selectively protects against obesity induced by diets high in saturated fat. Obesity (Silver Spring), 2008. 16(6): p. 1248-55. 31. Arnold, L., et al., Inflammatory monocytes recruited after skeletal muscle injury switch into antiinflammatory macrophages to support myogenesis. J Exp Med, 2007. 204(5): p. 1057-69. 32. Brigitte, M., et al., Muscle resident macrophages control the immune cell reaction in a mouse model of notexin-induced myoinjury. Arthritis Rheum, 2010. 62(1): p. 268-79. 33. Kim, T.W., et al., A critical role for IRAK4 kinase activity in Toll-like receptor-mediated innate immunity. J Exp Med, 2007. 204(5): p. 1025-36. 34. Li, X., IRAK4 in TLR/IL-1R signaling: possible clinical applications. Eur J Immunol, 2008. 38(3): p. 614-8. 35. Lerman, I., et al., Genetic variability in forced and voluntary endurance exercise performance in seven inbred mouse strains. J Appl Physiol (1985), 2002. 92(6): p. 2245- 55. 36. Laye, M.J., et al., Cessation of daily wheel running differentially alters fat oxidation capacity in liver, muscle, and adipose tissue. J Appl Physiol (1985), 2009. 106(1): p. 161- 8. 37. Ikeda, S., et al., Muscle type-specific response of PGC-1 alpha and oxidative enzymes during voluntary wheel running in mouse skeletal muscle. Acta Physiol (Oxf), 2006. 188(3-4): p. 217-23. 38. Nagatomo, F., et al., The effects of running exercise on oxidative capacity and PGC- 1alpha mRNA levels in the soleus muscle of rats with metabolic syndrome. J Physiol Sci, 2012. 62(2): p. 105-14. 39. Ropelle, E.R., et al., IL-6 and IL-10 anti-inflammatory activity links exercise to hypothalamic insulin and leptin sensitivity through IKKbeta and ER stress inhibition. PLoS Biol, 2010. 8(8). 40. Kelly, M., et al., Activation of AMP-activated protein kinase by interleukin-6 in rat skeletal muscle: association with changes in cAMP, energy state, and endogenous fuel mobilization. Diabetes, 2009. 58(9): p. 1953-60. 41. Krawczyk, C.M., et al., Toll-like receptor-induced changes in glycolytic metabolism regulate dendritic cell activation. Blood, 2010. 115(23): p. 4742-9. 42. Everts, B., et al., TLR-driven early glycolytic reprogramming via the kinases TBK1- IKKvarepsilon supports the anabolic demands of dendritic cell activation. Nat Immunol, 2014. 15(4): p. 323-32. 43. Ando, M., et al., Interleukin 6 enhances glycolysis through expression of the glycolytic enzymes hexokinase 2 and 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase-3. J Nippon Med Sch, 2010. 77(2): p. 97-105. 44. Bengt Saltin, P.D.G., Skeletal Muscle Adaptability: Significance for Metabolism and Performance. Compr Physiol 2011, 1983. Supplement 27: Handbook of Physiology, Skeletal Muscle(doi: 10.1002/cphy.cp100119): p. 555-631.

CHAPTER VI

Skeletal Muscle Toll-like Receptor-4 Deficiency Suppresses Exercise-induced Myokine Response and Diminishes Aerobic Training Adaptations in Mice.

Abstract

There is strong evidence supporting a crucial role for inflammatory cytokines in modulating

aerobic capacity and metabolic adaptations to exercise. Many reports have indicated that skeletal

muscle is the source of systemic elevations of cytokine levels, termed myokines, observed during

and post exercise. However, the exact mechanism mediating these muscle-derived inflammatory

cascades has yet to be elucidated. Pathogen-mediated activation of muscle TLR4 (mTLR4) is known to stimulate the production of pro-inflammatory myokines. However, it is not clear if mTLR4 is also implicated in the inflammatory responses associated with exercise and muscle contraction. As such, the purpose of this study was to investigate the role of m TLR4 signaling in modulating myokine production and release in active muscle and how mTLR4 deficiency

(mTLR4-/-) may impact the exercise-induced aerobic and oxidative adaptations. Intriguingly,

mTLR4 genetic inactivation depressed aerobic capacity improvements observed in WT mice

after treadmill training. Additionally, contrary to WT littermates, mTLT4-/- mice lacked treadmill

running-induced acute elevations in circulating myokines. In concert, mTLR4-/- mice also displayed attenuated myokine gene expression in response to in vivo electrical muscle stimulation (EMS) when compared with WT controls. Furthermore, mTLR4-/- mice experienced

metabolic limitations in skeletal muscle adaptability to exercise where they failed to match

oxidative improvements of WT littermates for glucose oxidation, fat oxidation, and

mitochondrial enzymatic capacity. Overall, these data disclose a direct association between

mTLR4 signaling and exercise-induced myokine production and release, a mechanism that may

account for metabolic maladaptation observed in mTLR4-/- mice. Furthermore, these data suggest

a link between mTLR4 and physiological determinants of maximal aerobic performance.

Key Words: skeletal muscle, toll-like receptor-4, treadmill training, myokines, aerobic capacity, metabolic adaptations

Introduction

Skeletal muscle is a major site controlling fuel depots and energy expenditure [1]. Exercise modulates whole body energy homeostasis via mechanisms manipulating skeletal muscle phenotype. The activation of exercise-responsive intracellular signaling pathways stimulates transcriptional events that dictate skeletal muscle properties. Inflammatory signaling plays a critical role in exerting the effect of exercise locally in the active muscle and systemically through endocrine mechanisms [2, 3]. Skeletal muscle has recently been recognized as a secretory organ [4]. Myokines are a class of inflammatory proteins produced by active muscle to convey many of the exercise-induced changes in muscle mass, contractile and metabolic properties [5].

Exercise-stimulated myokine production has been established to be an intensity- and duration-

dependent response [6, 7]. However, the precise molecular mechanism (s) modulating these

inflammatory cascades has yet to be elucidated. Post-exercise muscle glycogen content is

reported to be tightly related to levels of circulating cytokines and has been shown to regulate

myokine gene expression [8, 9]. Another line of evidence has suggested that exercise-induced

intracellular calcium flux and production of reactive oxygen species trigger transcriptional

factors promoting IL-6 synthesis [10, 11]. Nonetheless, the question remains as to whether the

exercise stress response follows similar inflammatory physiological pathways to those activated by other physical and pathogenic stimuli.

Due to a spontaneous genetic mutation in their endotoxin response locus, C3H/HeJ mice were used to initially identify the gene target for LPS [12]. This identification led to the discovery of

TLR4 which demonstrated a critical role in mediating the pro-inflammatory cytokine production in multiple tissues including skeletal muscle [13-15]. Intriguingly, in comparison to six other inbred mouse strains, C3H/HeJ mice showed the worst aerobic capacity as measured by maximal treadmill performance [16]. Moreover, contrary to respective WT controls, C3H/HeJ mice had a significant attenuation of exercise-induced IL-6 and IL-10 responses in the hypothalamus where this defect was found to abolish the exercise sensitization of insulin and leptin [17]. Together,

TLR4 is likely to exert its effect through modulating the exercise-associated cytokine response, a mechanism that may link TLR4 signaling to maximal exercise performance.

Exercise promotes the induction of metabolic and muscular injury by-products that invoke consequent inflammatory events. Depending on mode and duration, exercise has been documented to induce a transient increase in extracellular concentrations of damage-associated molecular patterns (DAMPs) and non-esterified fatty acids (NEFAs) in response to trauma and heightened energy demand, respectively [18, 19]. These molecules have been shown to stimulate

TLR4 signaling [20, 21]. A growing body of evidence indicates that TLR4 activation is vital for the resolution of muscle injury and long-term adaptations to chronic exercise [22, 23]. TLR4 has also been reported to modulate key cellular metabolic regulators including P38 MAPK and JNK

[23]. Nonetheless, the role TLR4 as a modulator of maximal exercise performance in response to aerobic training is still unanswered. Furthermore, the role of TLR4 in the exercise-induced cytokine response and its concomitant effects remains to be investigated. Skeletal muscle serves

as the primary tissue responsible for physical activity and systemic control of energy

homeostasis. As such, understanding the molecular network governing its contribution to

systemic adaptability to aerobic training is of great significance to the effectiveness of exercise

intervention programs.

Material and Methods

Animal husbandry and mTLR4-/- mice. Animal studies were performed under an approved

protocol by the Institutional Animal Care and Use Committee (IACUC) at Virginia Tech. The

muscle-specific TLR4 knockout (mTLR4-/-) mouse model on C57BL/6J background was created

at the Transgenic Core at the Pennington Biomedical Research Center using the Cre-mediated

recombination driven by the muscle creatine kinase promoter. Male mTLR4-/- and wild type

(WT) littermates were maintained on a 12-hour light/dark cycle and fed standard chow diet. All

experiments were performed following an overnight fast (10-12 hours), unless otherwise stated.

Acute and chronic treadmill running protocols. Forced treadmill running was used. Thirty

two mTLR4-/- and WT mice were randomized to either a sedentary control or exercise group.

Mice were acclimated to the treadmill for 2 days. Blood serum samples were collected and body

composition parameters were assessed prior (baseline) to and after (post-training) the completion of the exercise program. Due to limitations with mTLR4-/- mice available for the study, a

subgroup of 12 mice, out of the 32 mice, were used to examine changes in serum cytokine levels

in response to acute exhaustive treadmill running. The acute exercise protocol consists of one

bout of prolonged endurance exercise where mice ran until exhaustion at 0.30 m/sec. In addition

to the baseline serum samples that were collected at rest (the pre-exercise time point); another sample was collected immediately after the termination of the acute bout of treadmill running.

Allowing 3 days of recovery, mice then performed a maximal aerobic capacity test. Then mice were submitted to the treadmill training program which consists of a moderate intensity 50- minute run 5 days/week for 4 weeks. The protocol progressed into two phases. The first two weeks mice ran at 0.30 m/sec where the second two weeks were at 0.33 m/sec. After the completion of the treadmill running program mice performed a maximal aerobic capacity test.

Mice were sacrificed 36 hours after testing their aerobic capacity as a wash out period avoiding the effect of last bout exercise. Skeletal muscle (gastrocnemius, quadriceps, and soleus) were harvested for processing and data collection.

Figure 1. Schematic representation of the treadmill study design

Testing aerobic capacity. Using incremental treadmill running, the testing protocol begins with

5 minutes warm-up at low work rate followed by a stepwise increase in power output. The mice

went through incremental running starting at 0.20 m/sec and 0˚ inclination. The treadmill

velocity was increased by 0.03 m/s every two minutes until the mice reached exhaustion and

refused to continue running. Speed and total distance at which the mouse could not continue running were recorded. The testing session was to be terminated at any point at which a mouse met either of these exhaustion criteria:

1. Spending greater than 5 consecutive seconds on the shock grid without attempting to reengage the treadmill

2. Spending greater than 50% of its time on the shock grid

3. The third time a mouse was willing to sustain 2 seconds or more of shock without attempting to reengage the treadmill

In-vivo plantar flexor torque assay. In-vivo measures of plantar flexor torque (PFT) were obtained with mice under isoflurane anesthesia (platform at 35ºC). The mouse’s foot was secured to a torque transducer (Aurora Scientific) and the tibial nerve was stimulated at 2Hz for a series of contractions to fatigue (4 bouts X 8 minutes; 1 minute rest between bouts). At the end of the fatigue protocol, the leg was aseptically cleaned with povidone-iodine solution while the

mouse recovered from anesthesia (~2-5 minutes in 100% O2). Hindlimb PFT data were analyzed

using Dynamic Muscle Analysis (DMA) software versions 3.2 (Aurora Scientific).

Blood sampling and flow cytometry analysis. To determine serum cytokine profile, blood

samples were drawn using Microvette CB 300 Z tubes through the lateral saphenous vein which

runs dorsally and then laterally over the tarsal joint. Blood was then centrifuged at 8000 rpm for

10 minutes at 4oC for serum separation. Serum samples were blocked with mouse IL-6, IL-10,

MCP-1, IFN-ϒ, and TNF-α capture beads using the commercial Cytometric Bead Array (CBA)

Mouse Inflammation Kit (cat# 552364) according to the manufacturer's instructions (BD

Biosciences).

Skeletal muscle whole homogenate preparation for substrate metabolism. Approximately 50

mg of fresh muscle samples were immediately placed into 0.2 ml of a modified sucrose EDTA

medium (SET) on ice containing 250 mM sucrose, 1 mM EDTA, 10 mM tris-HCl, and 1mM

ATP, pH 7.4. Muscle samples were then minced with scissors and then SET buffer was added to

a 20-fold diluted (wt:vol) suspension. The minced samples were homogenized in a Potter-

Elvehjem glass homogenizer at 12 passes across 30 seconds at 150 rpm with a motor-driven

teflon pestle, and measures of fatty acid oxidation, glucose oxidation, pyruvate oxidation, and

maximal enzyme activities were performed.

Substrate metabolism. Red gastrocnemius and quadriceps muscles were collected for substrate

metabolism. End products of substrate oxidation as measured by [1-14C]-palmitic acid oxidation

to 14CO2 and [U-14C]-glucose oxidation to 14CO2 were assessed. For FAO and GO assays, tissue

homogenates from red skeletal muscles were incubated in 1μCi/mL of [1-14C]-palmitic acid or

1μCi/mL of [U-14C]-glucose respectively, for 1 hour and then acidified with 45% perchloric acid

to elute gaseous CO2. 14CO2 was trapped in 1M sodium hydroxide for 1 hour. The sodium

hydroxide was then sampled and placed in a liquid scintillation counter and the trapped 14CO2

was quantified.

Citrate synthase (CS) activity. Citrate synthase catalyzes the formation of citrate and CoASH

from acetyl-CoA and oxaloactetate. CoASH reduces DTNB and CS activity was determined

from the reduction of DTMB over time. Briefly, ten microliters of a 1:5 diluted muscle homogenate was added, in duplicate, to 170μl of a solution containing Tris buffer (0.1M, pH

8.3), DNTB (1mM, in 0.1M in Tris buffer) and oxaloacetate (0.01M, in 0.1M Tris buffer).

Following a 2min background reading, the spectrophotometer (SPECTRAmax ME, Molecular

Malate dehydrogenase (MDH) activity. MDH activity was measured spectrophotometrically at

340nm at 37°C. Briefly, 10ul of sample were pipetted in duplicate or triplicate in wells. Then,

290ul of reaction media (0.1 M potassium phosphate buffer, PH 7.4 plus 0.006 M oxaloacetic acid, prepared in potassium phosphate buffer plus 0.00375 M NADH, prepared in potassium phosphate buffer) was added to the wells and samples were read for 5 minutes at 340nm. The rate of disappearance of NADH was analyzed and expressed relative to protein content. MDH activity was calculated and reported as μmol/min/mg.

β-hydroxyacyl-CoA dehydrogenase (β-HAD) activity. For the determination of β-HAD activity, oxidation of NADH to NAD+ was measured. Volume of 35 μl of whole muscle homogenate was added to 190μl of a buffer containing 0.1M liquid triethanolamine, 5mM EDTA tetrasodium salt dihydrate, and 0.45mM NADH. The spectrophotometer (SPECTRAmax PLUS

384, Molecular Devices Corporation, Sunnyvale California) was calibrated and 15μl of 2mM acetoacetyl CoA was added to initiate the reaction. Absorbance was measured at 340 nm every

12 seconds for 5 minutes at 37C. β-HAD activity was calculated and reported as μmol/min/mg.

Cytochrome c oxidase (COX) activity. Cytochrome c oxidase enzyme activity was measured

spectrophotometrically at 550nm every 10 seconds for 7 minutes. Absorbance was measured

based on the oxidation of ferrocytochrome c to ferricytochrome c by cytochrome c oxidase. COX

activity was calculated and reported as μmol/min/mg.

RNA extraction and qRT-PCR. mRNA was extracted using an RNeasy Mini Kit (Qiagen,

Valencia, CA) and DNase I treatment (Qiagen, Valencia, CA), according to the manufacturer's

instructions. qRT-PCR was performed using an ABI PRISM 7900 Sequence Detection System

instrument and TaqMan Universal PCR Master Mix used according to the manufacturer's

specifications (Applied Biosystems, Foster City, CA). Target gene expression was normalized to

β-actin RNA levels. Primers and 5# FAM-labeled TaqMan probes were purchased as

prevalidated assays (Applied Biosystems, Foster City, CA).

Western blot analysis. Western blot analysis was performed using skeletal muscle homogenates

prepared in 50 mM Hepes, (pH7.5), 15 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 2 mM EDTA,

10% glycerol, 1 % Triton x-100, 10 mM NaP2O7, 100 mM NaFl, 10 mM PMSF, and 10 g/mL

aprotinin. Proteins (~30ug per sample) were separated using a 10% Criterion-Tris-HCl gel (Bio-

Rad, Hercules, CA) and subsequently transferred to PVDF membrane (Bio-Rad, Hercules, CA).

PVDF membrane was probed with primary antibody against GLUT4 (Cell Signaling, cat#

2213S) in a 1:1000 dilution.

Statistical analysis. Descriptive and inferential statistics methods were employed to analyze the collection of data of the study. The descriptive statistics as measured by mean and standard error were used to summarize the sample and to quantitatively describe the main features of the data

(e.g. body mass, lean mass, fat percent… etc.). Results are expressed as mean ± standard error of

the mean (SEM). As we were interested in studying the interaction effect of both exercise and

genotype differences (WT vs. mTLR4-/-) on manipulating exercise-induced myokine and

metabolic responses, the statistical significances of differences among control and experimental

groups were determined using the Two-Way ANOVA. If a significant F-ratio was obtained, comparisons between groups were made using Tukey’s post-hoc test. Furthermore, Student’s t- test was used to report the differences in gain scores between the control and experimental groups to better understand the genotype X exercise interaction effect. The level of significance was established as P < 0.05.

Results

Maximal aerobic capacity and body composition parameters. WT and mTLR4-/- littermates

experienced significant reductions in body fat percent (Figure 2A) in response to treadmill

training. Furthermore, data shows a main effect of the treadmill training protocol which

significantly increased running capacity of WT and mTLR4-/- mice, comparing between pre and

post-training protocol time points (Figure 2B). Data are presented as means ± SEM.

Figure 2. The effect of the treadmill training protocol on body fat and maximal aerobic capacity. Differences in body fat percent and aerobic capacity between WT and mTLR4-/- mice at pre and post-treadmill training time points. Treadmill training resulted in significant decreases in body fat percent (A) and increases in maximal running performance (B) in WT (n=8) and mTLR4-/-(n=8) mice. Data are presented as means ± SEM (# P < 0.05).

Force production in response to in-vivo electrical muscle stimulation. There was a significant main effect of time but no significant genotype X time interaction for the plantar flexor muscle torque (Figure 3). The force production profile showed a significant incremental decrease over

12 time points representing the time course of the EMS fatigue protocol. Data are presented as means ± SEM.

WT mTLR4-/-

Figure 3. Assessment of force production over the course of the in-vivo electrical muscle

stimulation protocol. The force production profile showed a significant incremental decrease

over 12 time points representing the 32- minute duration of the EMS fatigue protocol. WT (n=7) and mTLR4-/- mice (n=9) produced similar plantar flexor muscle torque. Data are presented as

means ± SEM (*P < 0.05).

Serum cytokine levels in response to acute exhaustive treadmill running. There was a

significant genotype X acute running interaction for circulating cytokines. IL-6 (4A), TNF-α

(4B), INF- γ (4C), but not MCP-1 (4D), levels in WT and TLR4-/- mice increased and decreased

respectively post-exercise. mTLR4-/- mice lacked normal cytokine responses following acute

exhaustive treadmill running. Data are presented as means ± SEM.

Figure 4. Assessment of inflammatory markers post-exhaustive treadmill running. In

response to acute exhaustive treadmill running , elevations of circulating IL-6 (A), TNF-α (B),

INF- γ (C), and MCP-1 (D) were suppressed in mTLR4-/- mice (n=6) compared with those

observed in WT littermates (n=6). Data are presented as means ± SEM (*P < 0.05).

Myokine gene expression in response to in-vivo electrical muscle stimulation. There was a

main effect of muscle stimulation and a significant genotype X muscle stimulation interaction for

mRNA levels of IL-6 and MCP-1. qRT-PCR analysis showed that in-vivo electrical stimulation

induced muscle contraction that resulted in significant increases in IL-6 (Figure 5A) and MCP-

1(Figure 5B) gene expression in WT and mTLR4-/- mice but to a significantly lesser extent in the

latter. Data are normalized to β-Actin and presented as means ± SEM.

Figure 5. Assessment of myokine gene expression in response to in-vivo electrical muscle

stimulation. Compared with respective augmentations found in WT controls (n=7), in-vivo

electrical muscle stimulation revealed a blunted contraction-induced mRNA increase of muscle

IL-6 (A) and MCP-1 (B) in mTLR4-/- mice (n=7). Data were normalized to β-Actin and presented as means ± SEM (*P < 0.05).

Metabolic adaptations in response to chronic treadmill training. Treadmill training resulted in significant increases in glucose oxidation in WT and mTLR4-/- mice (Figure 6A), but to a

significantly greater magnitude in WT mice (WT: +110% vs. mTLR4-/-: +43%: P<0.05) (Figure

6B). Although not statistically significant, there was a trend of an increase in levels of FAO in

WT mice, whereas mTLR4-/- littermates showed a completely blunted response to treadmill

training (Figure 6C). Furthermore, while WT mice witnessed trends of augmentations in enzymatic activities of CS (Figure 7A), MDH (Figure 7B), β-HAD (Figure 7C), and COX

(Figure 7D) in response to treadmill training, mTLR4-/- mice exhibited activity levels similar to

the basal rates observed in their untrained counterparts. Notably, each of the mTLR4 deletion

and treadmill training independently induced main effects manifested in significant

augmentations in β-HAD enzymatic activity (Figure 7C: P<0.05) Data are presented as means ±

SEM.

Figure 6. Assessment of changes in oxidative metabolism in response to treadmill training.

Compared with that shown in respective WT littermates (n=8), treadmill training revealed a significantly less pronounced increase in GO (A, B) in mTLR4-/- mice (n=8). However, FAO levels (C) remained unchanged in WT and mTLR4-/- mice. Data are presented as means ± SEM

(*P < 0.05).

Figure 7. Assessment of changes in mitochondrial maximal enzymatic activity in response

to treadmill training. Although not statistically significant, WT mice (n=8) showed trends of

increases in CS activity (A), MDH (B), β-HAD (C), and COX (D) in response to treadmill

training compared with a blunted response characterized in mTLR4-/- mice (n=8). However,

albeit mostly driven by WT mice, treadmill training induced a main effect of a significant

increase in β-HAD activity (C) regardless of the genotype differences. Meanwhile, mTLR4

deletion appears to have a main effect of a significant increase in β-HAD activity (C) compared

with WT littermates. Data are presented as means ± SEM (#P < 0.05 WT vs. mTLR4-/-,*P < 0.05

Sedentary vs. Trained).

Gene expression in response to chronic treadmill training. There was a significant genotype

X treadmill training interaction for GLUT4 mRNA levels (Figure 8A) and protein content

(Figure 9). Treadmill trained mTLR4-/- mice sustained the basal levels observed in respective

sedentary controls, whereas WT mice significantly increased GLUT4 expression in response to

treadmill training. No significant changes were observed in mRNA levels of PPAR-delta (Figure

8B), SOD2 (Figure 8C), IL-6 (Figure 8D), and MCP-1 (Figure 8E). Data are presented as means

± SEM.

Figure 8. Assessment of changes in gene expression of key metabolic and inflammatory genes in response to treadmill training. GLUT4 mRNA (A) increased significantly in treadmill trained WT mice (n=8), but not in trained mTLR4-/- (n=8), compared with respective sedentary

controls. PPAR-delta (B), SOD2 (C), IL-6 (D), and MCP-1 (E) mRNA remained unchanged.

Data are presented as means ± SEM (*P < 0.05).

Figure 9. Assessment of changes in GLUT4 protein levels in response to treadmill training.

GLUT4 protein content increased significantly in treadmill trained WT mice (n=8), but not in

trained mTLR4-/- (n=8), compared with respective sedentary controls. Representative western blots are included below their respective columns. Data are presented as means ± SEM (*P <

0.05).

Discussion

TLR4 is an exercise-responsive cellular receptor upstream from IL-6 and other inflammatory

cytokines that have demonstrated critical roles in exercise adaptability. Due to a mutation in the

TLR4 gene, C3H/HeJ mice are widely used in different research areas. Intriguingly, C3H/HeJ

mice were shown to lack the exercise-induced hypothalamic IL-6 response and the associated

insulin and leptin sensitization effect [17]. Meanwhile, Faldt and colleagues have reported

reduced treadmill endurance in IL-6 deficient mice [24]. Together, these findings gained our

impetus to explore the significance of TLR4 signaling to maximal exercise performance and its concomitant myokine and metabolic responses in skeletal muscle as a master regulator of whole body energy homeostasis. Towards this goal, we used an mTLR4-/- mouse model and pursued tightly standardized exercise and muscle stimulation modalities.

The main findings of this study were 1) Skeletal muscle TLR4 gene inactivation resulted in diminished muscle contraction-induced myokine gene expression as well as attenuation in the post-exercise elevations of circulating cytokines; and 2) mTLR4 signaling appears to play a critical role in modulating metabolic adaptations in response to aerobic training. 3) Maximal exercise performance is not a function of mTLR4 signaling. Designing an exercise experiment can be pursued by multiple approaches including voluntary and forced exercise modalities. The inflammatory and metabolic signaling associated with exercise performance has repeatedly been reported to be a mode- and duration-dependent response. With this in mind, treadmill running has best served our goal where we were able to control the intensity and amount of muscle work to limit the confounding factors that may arise from receiving different doses of exercise.

In addition to controlling the exercise dose, the diffusion characteristics of the muscle cell microenvironment are of fundamental importance in gene transcription regulation of skeletal muscle [25-28]. The extracellular milieu of a myocyte contains ions, metabolic substrates, neuroactive chemicals, signaling proteins and other substances that can profoundly alter the intramuscular signaling pathways [28-31]. At the same time, the extracellular space of the muscle is a highly complex medium that can be modulated by several factors including age [32], fat mass [33] and stress perception [34] each of which is unique to each experimental animal. As such, to minimize the mouse to mouse variability errors we also utilized in-vivo electrical muscle

stimulation to induce muscle contraction in one leg while the contralateral leg was used as a negative control.

The present study is the first to specifically examine the role of mTLR4 signaling in mediating the exercise-induced myokine response and how this may modulate aerobic capacity and skeletal muscle metabolism. Our data show no differences between WT and mTLR4-/- mice in basal

(before training) aerobic capacity. Our observations are consistent with those reported by H.

Zbinden and colleagues who did not detect significant differences in the maximal treadmill

running velocity between WT and transgenic TLR4-/- mice [23]. However, although not

statistically significant, our model shows that mTLR4 gene deletion resulted in a trend of

attenuation in the aerobic training-induced improvements in maximal treadmill performance

observed in the WT littermates. Noteworthy, mouse breeding and sample size issues may

generate experimental limitations impacting the power of the statistical inference tests employed

in data analysis.

Myokine response to exercise has always been shown to be mediated through muscle

contraction-stimulated intracellular signaling. Intramuscular exercise-sensitive components

including glycogen content, calcium flux, as well as reactive oxygen species have been

implicated in modulating myokine synthesis and release [9, 35]. Distinguishably, we propose

that the myokine cascades associated with exercise can also be modulated, in part, by systemic

factors (e.g. NEFA, HSP70) [36]that may exert their effects through cellular receptors localized

in the extracellular matrix including TLR4. Herein, we demonstrate that muscle-contraction

induced myokine gene expression entails mTLR4 signaling. As expected, mTLR4-/- failed to

activate IL-6 and MCP-1 gene transcription up to the levels observed in WT littermates in

response to in-vivo electrical muscle stimulation. This was further supported by the lack of

normal elevations in circulating cytokines in mTLR4-/- mice subjected to acute, exhaustive

treadmill running, unlike the increase seen in WT littermates. In concert, it was previously

reported that exercise-induced hypothalamic IL-6 and IL-10 responses were repressed in

C3H/HeJ mice that possess a dysfunctional TLR4 gene.

Impaired exercise-induced myokine response in mTLR4-/- mice was accompanied by marked

perturbations in metabolic adaptations to exercise training. Although treadmill training

contributed to an increase in levels of glucose oxidation in mTLR4-/- mice, yet this increase

failed to reach the magnitude of improvements achieved by WT. We have previously showed

that voluntary wheel running resulted in a substantial increase in upstream glycolytic

intermediates in WT mice, but not in mTLR4-/-, that may be facilitated, in part, by improved

glucose uptake. A major target for TLR4 and its downstream myokine signaling in skeletal

muscle, as an insulin-sensitive tissue, is GLUT4. There is a direct correlation between an

increase in insulin sensitivity and increased cell surface GLUT4 content. Exercise training results

in adaptations that facilitate insulin-mediated glucose uptake and are regulated by different

mechanisms. It leads to changes in gene expression, signaling, greater blood flow and changes in

GLUT4 protein exocytosis and endocytosis [37]. Our current data suggest a link between mTLR4 and GLUT4 gene reprogramming in response to exercise training. GLUT4 transcriptional adaptations following treadmill training, as measured by mRNA and protein levels, were diminished in mTLR4-/- mice compared to the significant augmentations achieved by

trained WT littermates. Furthermore, it has been reported that acute IL-6 exposure improves

insulin-stimulated glucose disposal in humans and glucose uptake in-vitro [38]. So the fact that mTLR4 gene disruption resulted in a depressed IL-6 response may explain the alterations observed in glucose trafficking and metabolism in mTLR4-/- mice [35, 38, 39]. Moreover in

response to treadmill training, WT mice had a trend of an increase in bet-oxidation which was accompanied by a trend of an increase in fatty acid utilization, though did not reach statistical significance. Collectively, these adjustments in carbohydrate metabolism in WT mice possibly increased the substrate supply to the TCA cycle but resulted only in trends of increases, with no statistical significance, in the mitochondrial oxidative enzymes citrate synthase, malate dehydrogenase, and cytochrome c oxidase, a marker for the ETC activity. This could be justified,

in part, by lack of statistical power due to small sample size and difficulties with matching the

different experimental groups. In contrast, mTLR4-/- mice exhibited a comprehensive inhibition

to aerobic-training induced metabolic adaptations. Indeed, TLR4 signaling was shown to be

essential to activate key regulators of cell energy homeostasis including MAPK [23].

Overall, our results demonstrate a critical role for mTLR4 in modulating the host adaptability to

aerobic training outcomes, specifically the metabolic responses. As such, further studies are of

great importance to explore a potential skeletal muscle fiber type–specific response to the

interplay between the muscle specific deletion of TLR4 and different modes of exercise. This

approach may lead to identifying an exercise modality that could bypass the genotype and

phonotypical perturbations in mTLR4 signaling which appear to impair aerobic training

outcomes in oxidative muscle. Furthermore, future studies should aim at characterizing

physiological adaptations to muscle work in the mTLR4-/- mouse model including muscle and cardiac functions and properties that modulate maximal aerobic capacity.

References

1. Zurlo, F., et al., Skeletal muscle metabolism is a major determinant of resting energy expenditure. J Clin Invest, 1990. 86(5): p. 1423-7. 2. Pedersen, B.K., Edward F. Adolph distinguished lecture: muscle as an endocrine organ: IL-6 and other myokines. J Appl Physiol (1985), 2009. 107(4): p. 1006-14. 3. Schnyder, S. and C. Handschin, Skeletal muscle as an endocrine organ: PGC-1alpha, myokines and exercise. Bone, 2015. 80: p. 115-25. 4. Pedersen, B.K. and M.A. Febbraio, Muscles, exercise and obesity: skeletal muscle as a secretory organ. Nat Rev Endocrinol, 2012. 8(8): p. 457-65. 5. Costamagna, D., et al., Role of Inflammation in Muscle Homeostasis and Myogenesis. Mediators Inflamm, 2015. 2015: p. 805172. 6. Peake, J.M., et al., Plasma cytokine changes in relation to exercise intensity and muscle damage. Eur J Appl Physiol, 2005. 95(5-6): p. 514-21. 7. Pedersen, B.K., Special feature for the Olympics: effects of exercise on the immune system: exercise and cytokines. Immunol Cell Biol, 2000. 78(5): p. 532-5. 8. Keller, C., et al., Effect of exercise, training, and glycogen availability on IL-6 receptor expression in human skeletal muscle. Journal of Applied Physiology, 2005. 99(6): p. 2075-2079. 9. Nieman, D.C., et al., Post-Exercise Skeletal Muscle Glycogen Related to Plasma Cytokines and Muscle IL-6 Protein Content, but not Muscle Cytokine mRNA Expression. Front Nutr, 2015. 2: p. 27. 10. Fischer, C.P., Interleukin-6 in acute exercise and training: what is the biological relevance. Exerc Immunol Rev, 2006. 12(6-33): p. 41. 11. Keller, C., et al. Human muscle cells express IL-6 via a Ca2+-dependent pathway. in JOURNAL OF PHYSIOLOGY-LONDON. 2002. CAMBRIDGE UNIV PRESS 40 WEST 20TH ST, NEW YORK, NY 10011-4221 USA. 12. Poltorak, A., et al., Defective LPS signaling in C3H/HeJ and C57BL/10ScCr mice: mutations in Tlr4 gene. Science, 1998. 282(5396): p. 2085-8. 13. Frost, R.A., G.J. Nystrom, and C.H. Lang, Lipopolysaccharide regulates proinflammatory cytokine expression in mouse myoblasts and skeletal muscle. Am J Physiol Regul Integr Comp Physiol, 2002. 283(3): p. R698-709. 14. Song, M.J., et al., Activation of Toll-like receptor 4 is associated with insulin resistance in adipocytes. Biochem Biophys Res Commun, 2006. 346(3): p. 739-45. 15. Gustot, T., et al., Differential liver sensitization to toll-like receptor pathways in mice with alcoholic fatty liver. Hepatology, 2006. 43(5): p. 989-1000. 16. Lerman, I., et al., Genetic variability in forced and voluntary endurance exercise performance in seven inbred mouse strains. J Appl Physiol (1985), 2002. 92(6): p. 2245- 55. 17. Ropelle, E.R., et al., IL-6 and IL-10 anti-inflammatory activity links exercise to hypothalamic insulin and leptin sensitivity through IKKbeta and ER stress inhibition. PLoS Biol, 2010. 8(8). 18. Seys, S.F., et al., Damage-associated molecular pattern and innate cytokine release in the airways of competitive swimmers. Allergy, 2015. 70(2): p. 187-94.

19. Walker, M., et al., Role of plasma non-esterified fatty acids during and after exercise. Clin Sci (Lond), 1991. 81(3): p. 319-25. 20. Shi, H., et al., TLR4 links innate immunity and fatty acid-induced insulin resistance. J Clin Invest, 2006. 116(11): p. 3015-25. 21. Mencin, A., J. Kluwe, and R.F. Schwabe, Toll-like receptors as targets in chronic liver diseases. Gut, 2009. 58(5): p. 704-20. 22. Paiva-Oliveira, E.L., et al., TLR4 signaling protects from excessive muscular damage induced by Bothrops jararacussu snake venom. Toxicon, 2012. 60(8): p. 1396-403. 23. Zbinden-Foncea, H., et al., TLR2 and TLR4 activate p38 MAPK and JNK during endurance exercise in skeletal muscle. Med Sci Sports Exerc, 2012. 44(8): p. 1463-72. 24. Faldt, J., et al., Reduced exercise endurance in interleukin-6-deficient mice. Endocrinology, 2004. 145(6): p. 2680-6. 25. Lauritzen, H.P., et al., Imaging of insulin signaling in skeletal muscle of living mice shows major role of T-tubules. Diabetes, 2006. 55(5): p. 1300-6. 26. Shorten, P.R., C.D. McMahon, and T.K. Soboleva, Insulin transport within skeletal muscle transverse tubule networks. Biophys J, 2007. 93(9): p. 3001-7. 27. Robinson, R., Fast, slow, or not at all: the extracellular matrix controls movement of signaling molecules. PLoS Biol, 2012. 10(7): p. e1001362. 28. Muller, P. and A.F. Schier, Extracellular movement of signaling molecules. Dev Cell, 2011. 21(1): p. 145-58. 29. Brum, G., E. Rios, and E. Stefani, Effects of extracellular calcium on calcium movements of excitation-contraction coupling in frog skeletal muscle fibres. J Physiol, 1988. 398: p. 441-73. 30. Huang, C.F. and J. Schmidt, Calcium influx blocks the skeletal muscle acetylcholine receptor alpha-subunit gene in vivo. FEBS Lett, 1994. 338(3): p. 277-80. 31. Berchtold, M.W., H. Brinkmeier, and M. Muntener, Calcium ion in skeletal muscle: its crucial role for muscle function, plasticity, and disease. Physiol Rev, 2000. 80(3): p. 1215-65. 32. Yamada, Y., et al., Extracellular water may mask actual muscle atrophy during aging. J Gerontol A Biol Sci Med Sci, 2010. 65(5): p. 510-6. 33. Sinha, R., et al., Assessment of skeletal muscle triglyceride content by (1)H nuclear magnetic resonance spectroscopy in lean and obese adolescents: relationships to insulin sensitivity, total body fat, and central adiposity. Diabetes, 2002. 51(4): p. 1022-7. 34. Edeleva, E.V. and H.R. Shcherbata, Stress-induced ECM alteration modulates cellular microRNAs that feedback to readjust the extracellular environment and cell behavior. Front Genet, 2013. 4: p. 305. 35. Kosmidou, I., et al., Production of interleukin-6 by skeletal myotubes: role of reactive oxygen species. Am J Respir Cell Mol Biol, 2002. 26(5): p. 587-93. 36. Heck, T.G., C.M. Scholer, and P.I. de Bittencourt, HSP70 expression: does it a novel fatigue signalling factor from immune system to the brain? Cell Biochem Funct, 2011. 29(3): p. 215-26. 37. Lehnen AM, A.K., Markoski MM, Schaan BDA Changes in the GLUT4 Expression by Acute Exercise, Exercise Training and Detraining in Experimental Models. J Diabetes Metab S10:002., 2012.

38. Carey, A.L., et al., Interleukin-6 increases insulin-stimulated glucose disposal in humans and glucose uptake and fatty acid oxidation in vitro via AMP-activated protein kinase. Diabetes, 2006. 55(10): p. 2688-97. 39. Glund, S., et al., Interleukin-6 directly increases glucose metabolism in resting human skeletal muscle. Diabetes, 2007. 56(6): p. 1630-7.

CHAPTER VII

Conclusions and Future Directions

TLR4 plays a critical role in skeletal muscle in both health and disease. The understanding of

TLR4 signaling in skeletal muscle has been limited by the lack of a muscle-specific manipulation

that enables the study of the functional signaling of muscle TLR4 in vivo. Our novel mouse

model with the muscle-specific deletion of TLR4 bridges this research gap and methodological

limitation. Our in vivo investigations suggest an essential function for muscle TLR4 in

developing some of the health benefits commonly reported with exercise interventions. Findings

from the current study imply that muscle TLR4 mediates the inflammatory cascades associated

with exercise, thereby modulating the concomitant metabolic adaptations in oxidative tissues.

TLR4 appears to control exercise-mediated oxidative metabolism by initiating signals that secure

continuous supply of metabolic substrates to the working muscle as well as boosting the

enzymatic machinery serving the associated metabolic pathways. These effects could impact

lifestyle-related diseases.

Developed countries with a westernized diet and sedentary lifestyle suffer from chronic ailments

with inflammatory etiology, including obesity and type-2 diabetes, which may relate to

inappropriate TLR4 signaling. In these conditions, the power of TLR4 is often modulated by

negative regulators suppressing the chronic excess of inflammatory signaling. These negative effects may compromise TLR4 signaling and abolish its mediatory role in exercise-induced metabolic adaptations and any consequent health benefits. In obese humans, TLR4 signaling is chronically activated in a pathological manner associated with a concurrent increase in the expression of suppressors of cytokine signaling (SOCS) proteins in metabolic tissues. It has been

shown that elevations in the expression of SOCSs mediate inhibitory effects on insulin signaling

and glucose metabolism. In this context, the impaired inflammatory tone with chronic anti- inflammatory signaling cascades could potentially disrupt normal TLR4-associated inflammatory and metabolic mechanisms. Further studies are needed to characterize the differential physiological and metabolic response of obese and healthy lean individuals to different exercise modalities. This may delineate the adaptive processes and mechanisms that are potentially altered, which may lead to the identification of key molecular targets for therapeutic interventions.

APPENDICES

Appendix A: Generation of Skeletal Muscle-Specific TLR4 Knockout Mice

A targeting vector was generated using recombineering. Isogenic DNA containing TLR4 gene

was retrieved from a clone (RP24-209H15) of the C57BL6 BAC genomic library via gap repair.

The loxP-neo-loxP was inserted into intron 1 and the third loxP site into intron 2 via homologous

recombination to make a targeting vector with the exon 2 flanked by the last two loxP sites. For

mouse ES cell targeting, NotI-linearized TLR4 targeting vector DNA, consisting of 2 kb 5′-arm and 8 kb 3′-arm, was electroporated into C57BL/6J ES cell line Bruce 4 (a gift from the NCI,

NIH). Correct targeting was confirmed using high fidelity PCR analyses and DNA sequencing.

ES cells were injected into the blastocysts of female albino B6 strain C57BL/6J-Tyrc-2J obtained

from The Jackson Laboratory. Chimeras were then crossed with the albino B6 strain to screen for

heterozygous carriers of the modified allele. The Pgk-neo gene was removed by mating heterozygotes (TLR4loxneo/+) with B6.FVB-Tg (EIIa-cre) to generate heterozygous mice

(TLR4lox/+). To generate muscle-specific KO mice for TLR4, TLR4lox/+ mice were bred to

MCK-Cre recombinase mice (from Gerald Bothe, Wadsworth Center, Albany, NY) that were backcrossed for 10 generations to C57BL6 mice. Therefore, all analyses are from mice on the

C57BL6 background.