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Equine Skeletal Muscle Mitochondrial Function and Regeneration Capacity with Aging

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Title:
Equine Skeletal Muscle Mitochondrial Function and Regeneration Capacity with Aging
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Li, Chengcheng
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[Gainesville, Fla.]
Florida
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University of Florida
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english
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Doctorate ( Ph.D.)
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University of Florida
Degree Disciplines:
Animal Molecular and Cellular Biology
Committee Chair:
WOHLGEMUTH,STEPHANIE
Committee Co-Chair:
WARREN,LORI KAY
Committee Members:
DRIVER,JOHN P
LEEUWENBURGH,CHRISTIAAN

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Subjects / Keywords:
aging -- mitochondria
Animal Molecular and Cellular Biology -- Dissertations, Academic -- UF
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bibliography ( marcgt )
theses ( marcgt )
government publication (state, provincial, terriorial, dependent) ( marcgt )
born-digital ( sobekcm )
Electronic Thesis or Dissertation
Animal Molecular and Cellular Biology thesis, Ph.D.

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Abstract:
Skeletal muscle aerobic capacity, mitochondrial function and regenerative capacity have been found to decline with age in humans and rodents. However, not much is known about age-related changes in mitochondrial function in equine skeletal muscle. The objective of this dissertation was to: 1) evaluate differences in skeletal muscle mitochodnrial function between young and aged horses; 2) address underlying mechanisms for the age-related alterations in mitochondrial function; and 3) examine differences in the intrinsic regenerative capacity of muscle stem cells isolated from skeletal muscle of young and aged horses. We first compared muscle aerobic capacity, especially mitochodnrial density and function in gluteus medius (GLU) and triceps brachii (TRI) muscle between young and aged American Quarter Horses. Equine skeletal muscle aging was accompanied by a shift in fiber type composition towards a higher percentage of type I and IIA muscle fibers, decrease in mitochondrial density and cytochrome c oxidase activity, but preserved mitochodnrial respiratory function. To further understand the underlying cause for the age-associated decrement of mitochondrial density and function in equine skeletal muscle, expression of factors involved in mitochondria biogenesis and mitochondria-selective autophagy pathways, two of the most prominent quality control mechanisms that have been described, were analyzed. Our data suggest that mitochondrial content and biogenesis markers were reduced in aged TRI, and that autophagic activity was impaired in both muscles with age, albeit more pronounced in the TRI muscle. Myogenic stem cells, commonly referred to as satellite cells, are responsible for muscle growth and repair in adults. However, the ability to regenerate muscle and replace damaged myofibers declines with age. We then asked whether the intrinsic changes within an aged satellite cell would cause alteration in regenerative capacity in equine. In vitro studies with primary satellite cell cultures under standard conditions suggest that satellite cells isolated from aged horses displayed compromised proliferative, differentiation and fusion capacity in vitro. In line with compromised myogenic potential of aged muscle-derived satellite cells, there were age-related alterations in mitochondrial biogenesis and autophagy pathways, with satellite cells derived from TRI being more susceptible to impairments with age. ( en )
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In the series University of Florida Digital Collections.
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Includes vita.
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This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Thesis:
Thesis (Ph.D.)--University of Florida, 2017.
Local:
Adviser: WOHLGEMUTH,STEPHANIE.
Local:
Co-adviser: WARREN,LORI KAY.
Electronic Access:
RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2018-02-28
Statement of Responsibility:
by Chengcheng Li.

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EQUINE SKELETAL MUSCLE MITOCHONDRIAL FUNCTION AND REGENERATION CAPACITY WITH AGING By CHENGCHENG LI A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2017

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2017 Chengcheng Li

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To my family

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4 ACKNOWLEDGMENTS I would like to thank numerous wonderful people who have supported me throughout my PhD life I must thank my advisor Dr. Stephanie Wohlgemuth, a great scientist I will never forget. She is very knowledgeable and has taught me many great skills regarding to muscle physiology. Without her excellent guidance and dedication, I would never complete m y dissertation. I want to give Dr. Wohlgemuth one more thanks for giving me her friendship and life advice. She has been taking care of me through all these years. Moreover, she has never forgotten to prepare cute presents for my little girl. I would also like to thank the rest of my graduate committee Dr. Lori Warren, Dr. John Driver and Dr. Christiaan Leeuwenburgh for their invaluable academic support. Dr. Warren has been so helpful with planning and designing all the horse studies, and guided me through the experiments. Dr. Driver and Dr. Leeuwenburgh have made themselves so approachable to help me to develop my background in physiology and molecular biology. There are also so many faculty members that have made me complete my projects. I thank Dr. John Bromfield, Dr. Corwin Nelson, Dr. Peter Hansen and Dr. Timothy Hackman for letting me use their equipment. I thank Dr. Tracy Scheffler and Dr. Jason Scheffler for helping me troubleshooting experimental issues. Special thanks goes to Dr. David Julian from Department of Biology, who was willing to train me how to use fluorescent equipment. W arm thanks to Ted Broome for devoting so much time to help me with develop ing experimential protocol s Additionally, I also would like to express my appreciation to all graduate students that helped me with sample collection and generously gave me support through my life at UF. I thank a former graduate student Sarah White, who is currently an assistant pro fessor of equine physiology at

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5 Texas A&M University. Sarah has taught me muscle biopsy skills that I could never imagine I can do on my own now. To my lab mate Ariane Sousa: thank you for donating your time to assist with sample collection. Thank Paula Tri bulo, Vernica Negrn Prez, Sofia Ortega and numerous graduate students who shared ideas and gave advice during my program. I also would like to thank the staff, including Justin Callaham and Sciences Center and Horse Teaching Unit, for assistance. Lastly, my warmest thanks must be to my family. I must thank my parents and my parents in law for making my life sweet and wonderful. Thank them for coming to take care of my daughter to make me co ncentrate on my research. Their love and support made me who I am today. To my beloved little girl, Emilia: you are the best daughter I could ever have, and your smiles encourage me. I also thank my dear husband, Peng Liu, for his continued love. I conside r myself to be truly fortunate to have him stand by to support and encourage me during the challenges of graduate program to make the completion of my dissertation possible.

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6 TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................ ................................ ................................ .. 4 LIST OF TABLES ................................ ................................ ................................ ............ 9 LIST OF FIGURES ................................ ................................ ................................ ........ 10 LIST OF ABBR EVIATIONS ................................ ................................ ........................... 12 ABSTRACT ................................ ................................ ................................ ................... 14 CHAPTER 1 LITERATURE REVIEW ................................ ................................ .......................... 16 I ntroduction ................................ ................................ ................................ ............. 16 Aging Related Decline in Skeletal Muscle Mass and Function ............................... 18 Ch anges in Skeletal Muscle Fiber Type Composition with Aging ..................... 20 Changes in Muscle Energy Metabolism with Aging ................................ .......... 24 Changes in Mitochondrial Function with Aging ................................ ................. 28 Mitochondrial Quality Control Mechanisms ................................ ............................. 32 Mitochondrial Proteases and Chaperones ................................ ........................ 33 Fission and Fusion ................................ ................................ ........................... 33 Autophagy ................................ ................................ ................................ ........ 34 Mitochondrial Biogenesis ................................ ................................ .................. 36 Crosstalk between Different Mitochondrial Quality Control Mechanisms ......... 37 Aging and Muscle Regeneration Capacity ................................ .............................. 39 Satellite Cells and Myogenesis ................................ ................................ ......... 39 Aging Related Decline in Satellite Cell Function ................................ ............... 42 2 EFFECTS OF AGING ON MITOCHONDRIAL FUNCTION IN SKELETAL MUSCLE OF AMERICAN QUARTER HORSES ................................ .................... 47 Background ................................ ................................ ................................ ............. 47 Materials and Methods ................................ ................................ ............................ 51 Animals ................................ ................................ ................................ ............. 51 Skeletal Muscle Sampling ................................ ................................ ................ 51 Preparation of Permeabilized Muscle Fibers ................................ .................... 52 High Resolution Respirometry ................................ ................................ .......... 53 Sample Preparation and SDS PAGE for Myosin Heavy Chain Analysis .......... 54 Spectrophotometric Determination of Enzyme Activities ................................ .. 55 Statistical Analyses ................................ ................................ .......................... 56 Results ................................ ................................ ................................ .................... 56 Aging and Muscle Fiber Type Composition ................................ ...................... 56 Effect of Age on Mitochondrial Density and Enzyme Activity ............................ 57

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7 Effects of Age on Mitochondrial Respiration ................................ ..................... 58 Effect of Aging on Coupling Control Ratios ................................ ...................... 59 Discussion ................................ ................................ ................................ .............. 60 3 SKELETAL MUSCLE FROM AGED AMERICAN QUARTER HORSES SHOWS IMPAIRMENTS IN MITOCHONDRIAL BIOGENESIS AND AUTOPHAGY ............ 83 Background ................................ ................................ ................................ ............. 83 Materials and Methods ................................ ................................ ............................ 85 Animals ................................ ................................ ................................ ............. 85 Muscle Tissue Sampling ................................ ................................ ................... 86 Analys is of MtDNA Copy Number ................................ ................................ .... 86 RNA Isolation ................................ ................................ ................................ ... 87 Analysis of mRNA Expression ................................ ................................ .......... 87 Analysis of Protein Expression by Western Blot ................................ ............... 88 Statistical Analysis ................................ ................................ ............................ 89 Results ................................ ................................ ................................ .................... 89 Mitochondrial Conten t Was Decreased in Aged Skeletal Muscle ..................... 89 Mitochondrial Biogenesis Was Impaired with Age ................................ ............ 90 Transcript Level of MtDNA Encoded Genes Was Not Affected by Age ............ 91 Autophagic Capacity Was Impaired with Age ................................ ................... 91 Autophagosome Formation Was Impacted by Age ................................ .......... 92 Transcript Level of Lysosomal Degradation Marker LAMP2 Was Not Impacted by Age ................................ ................................ ........................... 93 Discussion ................................ ................................ ................................ .............. 93 4 AGE RELATED CHAN GES IN MYOGENIC CAPACITY OF SATELLITE CELLS OBTAINED FROM AMERICAN QUARTER HORSES ................................ .......... 112 Background ................................ ................................ ................................ ........... 112 Materials and Methods ................................ ................................ .......................... 115 Animals and Muscle Sample Collection ................................ .......................... 115 Satellite Cell Isolation ................................ ................................ ..................... 116 Proliferation and Differentiation Assays ................................ .......................... 116 Measurement of Myoblast Differentiation and Fusion ................................ .... 117 Isolation of Total RNA and Real Time qPCR ................................ .................. 118 MtDNA Copy Number Measurement ................................ .............................. 119 Immunoblotting ................................ ................................ ............................... 120 Statistical Analysis ................................ ................................ .......................... 121 Results ................................ ................................ ................................ .................. 121 Satellite Cells from Aged Horses Showed Reduced Myogenic Potential ....... 121 Mitochondrial DNA Copy Was Elevated with Age ................................ .......... 122 Mitochondrial Genes Were Downregulated i n Differentiated Cells from Aged TRI Muscle ................................ ................................ ................................ .. 123 Satellite Cells Derived from Aged Muscle Exhibited Impaired Mitochondrial Quality Control Mechanism ................................ ................................ ......... 124 Discussion ................................ ................................ ................................ ............ 125

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8 5 CONCLUSIONS AND FUTURE DIRECTIONS ................................ .................... 144 APPENDIX A CITRATE SYNTHASE ACTIVITY PROTOCOL ................................ .................... 147 B CYTOCHROME C OXIDASE ACTIVITY PROTOCOL ................................ ......... 150 C 3 OH ACYL COA DEHYDROGENASE ACTIVITY PROTOCOL .......................... 152 D MYOSIN HEAVY CHAIN ISOFORMS IDENTIFICATION PROTOCOL ................ 155 E PERMEABILIZED FIBERS PROTOCOL ................................ .............................. 15 8 LIST OF REFERENCES ................................ ................................ ............................. 162 BIOGRAPHICAL SKETCH ................................ ................................ .......................... 195

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9 LIST OF TABLES Table page 2 1 Fiber type composition of gluteus medius and triceps brachii from American Quarter Horses ................................ ................................ ................................ ... 75 2 2 Effect of age on fiber type composition in equine skeletal muscle ...................... 76 3 1 Primers used for gene amplification in quantitative reverse transcription polymerase chain reaction ................................ ................................ ................ 103 D 1 SDS PAGE gel mixture ................................ ................................ .................... 155 E 1 BIOPS buffer, total volume = 1 L ................................ ................................ ...... 158 E 2 MiR05 buffer, total volume = 1 L ................................ ................................ ....... 158 E 3 Commonly used SUIT chemicals ................................ ................................ ...... 160 E 4 SUIT protocol used in C hapter 2 ................................ ................................ ...... 160

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10 LIST OF FIGURES Figure page 2 1 Respirometric protocol with permeabilized fibers from American Quarter Horse gluteus medius muscle ................................ ................................ ............. 77 2 2 Representative SDS polyacrylamide gel stained with Coomassie blue following electrophoretic separation ................................ ................................ ... 78 2 3 Enzyme activities in muscle tissue homogenat es from American Quarter Horses ................................ ................................ ................................ ................ 79 2 4 Mitochondrial respiration of p ermeabilized skeletal muscle fibers from American Quarter Horses ................................ ................................ ................... 80 2 5 Mitochondrial coupling control ratios of permeabilized skeletal muscle fibers from American Quarter Horses ................................ ................................ ........... 82 3 1 Protein expression of citrate synthase in skeletal muscle from America n Quarter Horses ................................ ................................ ................................ 104 3 2 Mitochondrial DNA copy number and transcript levels of factors associated with mitochondrial biogenesis in skeletal muscle from American Quarter Horses ................................ ................................ ................................ .............. 105 3 3 Transcript level of mtDNA encoded genes in skeletal muscle from American Q uarter Horses ................................ ................................ ................................ 107 3 4 Protein expression of p62 in skeletal musc le from American Quarter Horses .. 108 3 5 Gene and protein expression of autophagy regulatory proteins in skeletal muscle from American Quarter Horses ................................ ............................ 109 3 6 Transcript level of the LAMP2 gene in skeletal muscle from American Quarter Horses ................................ ................................ ................................ 111 4 1 Proliferation rate of satellite cells isolated from skelatlal muscle of American Quarter Horses ................................ ................................ ................................ 132 4 2 Expresion of myogenin in satellite cells isolated from skeletal muscle of American Quarter Horses ................................ ................................ ................. 133 4 3 Fusion capacity of satellite cells isolated from skeletal muscle of American Quarter Ho rses ................................ ................................ ................................ 134 4 4 The protein expression of citrate synthase in satellite cells isolated from skeletal muscle of American Quarter Horses ................................ ................... 135

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11 4 5 Mitochondrial DNA copy number in satellite cells isolated from skeletal muscle of American Quarter Horses ................................ ................................ 136 4 6 Transcript levels of mtDNA encoded genes in differentiated satellite cells ....... 137 4 7 Protein expression of Hsp60 in satellite cells during differentiation in vitro ....... 138 4 8 Transcript level of genes relevant to mitochondrial biogenesis in satellite cells during differentiation in vitro ................................ ................................ ............. 139 4 9 Protein expression of autophagy regulators in satellite cells during differentiation in vitro ................................ ................................ ........................ 140 4 10 Gene expression of LC3 in satellite cells during differentiation in vitro ............. 142 4 11 Transcript level of the LAMP2 gene in satellite cell isolated from skeletal muscle from American Quarter Horses ................................ ............................ 143

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12 LIST OF ABBREVIATIONS ADP Adenosine diphosphate ANT Adenine nucleotide transporter ATP Adenosine triphosphate COX Cytochrome c oxidase CS Citrate synthase ER Endoplasmic reticulum ETS Electron transport system FADH 2 Flavin adenine dinucleotide FGF Fibroblast growth factors 3 HADH 3 OH acyl CoA dehydrogenase HGF Hepatocyte growth factor LC3 Microtubule associated proteins 1 A/B light chain 3 MRF Myogenic regulatory factor MyHC Myosin heavy chain NADH Nicotinamide adenine dinucleotide OXPHOS Oxidative phosphorylation Pax3 Paired box protein 3 Pax7 Paired box protein 7 PCr Phosphocreatine PGC Peroxisome proliferator activated receptor ROS Reactive oxygen species SDS PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis TCA Tricarboxylic acid TFAM Mitochondrial transcription factor A

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13 TGF Transforming growth factor beta 1 UCP Uncoupling protein

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14 Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy EQUINE SKELETAL MUSCLE MITOCHONDRIAL FUNCTION AND REGENERATION CAPACITY WITH AGING By Chengcheng Li August 2017 Chair: Stephanie E. Wohlgemuth Major: Animal Molecular and Cellular Biology Skeletal muscle aero bic capacity, mitochondrial function and regenerative capacity have been found to decline with age in humans and rodents. However, not much is known about age related changes in mitochondrial func tion in equine skeletal muscle. The objective s of this dis sertation w ere to: 1) evaluate differences in skeletal muscl e mitocho nd rial function between young and aged horses; 2) address underlying mechanisms for t he age related alterations in mitochondrial function; and 3) examine differences in the intrinsic regenerative capacity of muscle stem cells isolated from skeletal muscle of young and aged horses. W e first compared muscle aerobic capacity, especially mitochodnrial density and function in gluteus medius (GLU) and triceps brachii (TRI) muscle between young and aged American Quarter Horses. Equine skeletal muscle aging was accompanied by a shift in fiber type composition towards a hi gher percentage of type I and IIA muscle fibers decrease in mitochondrial density and cytochrome c oxidase activity, but preserved mitochodnrial respiratory function.

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15 To further understand the underlying cause for the age associated decrement of mitochondrial density and function in equine skeletal muscle, expression of factor s involved in mitochondria l biogenesis and mitochondria l selective autophagy pathways two of the most prominent quality control mechanisms that have been described, were ana lyzed. Our data suggest that mitochondrial content and biogenesis markers were reduced in aged TRI and that autophagic activity was impaired in both muscles with age, a lbeit more pronounced in the TRI muscle. Myogenic stem cells, commonly referred to as s atellite cells, are responsible for muscle growth and repair after birth However, the ability to regenerate muscle and replace damaged myofibers declines with age We then asked whether the intrinsic changes within an aged satellite cell would cause alteration in regenerative capacity in equine s In vitro studies with primary satellite cell cultures under standard conditions suggest that s atellite cells isolated from aged horses display ed compromised proliferative, differentiation and fusion capacity in vitro In line with compromised myogenic potential of aged muscle derived satellite cells, there were age related alterations in mitochondrial biogenesis and autophagy pathways with satellite cells derived from TRI muscle being more susceptible to impa irments with age

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16 CHAPTER 1 LITERATURE REVIEW I ntroduction Biological aging is a natural phenomenon that occurs in every living species. With age, tissues and organs begin to lose their ability to function correctly, or to function at all. However, the rate of aging can vary between individuals, depending on fitness levels and environment al factors. Furthermore, not all tissues experience aging at the same rate and to the same extent. Some system s may change slowly and exhibit minor dysfunctions, while others show dramatic decline and impairment to a greater extent. The exact cause of aging is not completely understood yet. The theories of aging are many, and one of the most popular explanations is that as ce lls age they gradually become unable to get rid of the wastes and toxins, and eventually, they are no longer able to function properly. For this reason, post mitotic tissues such as heart, skeletal muscle, and brain are more likely to be affected by aging (Kwong & Sohal, 2000). Cells within these tissues are not or only to a minor extent mitotically active, and therefore rarely or not at all replaced. Together with increasingly insufficient cellular quality control, damage to macromolecules accumulates, lea ding to cellular dysfunction. Skeletal muscle is profoundly affected by aging, and its functional decline is characterized by a progressive loss of muscle mass and strength ( Delmonico et al. 2009). Previous studies in humans have shown that leg muscle mass was reduced by ~30% (Janssen et al. 2000), and that thigh muscle cross sectional area declined by ~25% with aging (Klitgaard et al. 1990). In line with the age related decline in thigh muscle mas s, the overall physical function was also decreased (Buford et al. 2012). Similarly, muscle torque, an indicator of muscle quality or function, was 30 40% lower in

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17 sedentary elderly compared to young subjects (Klitgaard et al. 1990). The gradual loss of muscle mass is also linked to a progressive reduction in the regenerative capacity of skeletal muscle. Regeneration of adult skeletal muscle mainly depends on muscle stem cells, also known as satellite cells, located between the basal lamina and muscle fib er membrane (Mauro, 1961). Under normal conditions, satellite cells are rapidly activated by muscle damage and other stimuli (such as exercise), and mediate quick and complete repair or growth by either forming new fibers or fusing with existing fibers. Wi th age, however, skeletal muscle regenerativ e capacity declines (Carlson & Conboy, 2007), which can be partially explained by a decline in satellite cell function, as reported for mice ( Conboy et al. 2005; Chakkalakal et al. 2012 ; ) and humans ( Roth et al. 2000 b ). For this reason, repair or replacement of damaged muscle fibers is attenuated in elderly individuals This impairment in regenerative and growth capacity cont ributes to the decline in muscle mass and strength observed with aging (Jang et al. 2011). Although numerous factors have been implicated in the age associated decline in skeletal muscle mass, function, and repair capacity dysfunctional mitochondria in p articular are thought to play a primary role in this process. The mitochondria are the main generators of cellular energy. At the same time, they produce and release free radicals, which can cause damage to mitochondria themselves as well as other cellular constituents. For this reason, the mitochondrial theory of aging proposes that the progressive accumulation of damage to mitochondria is the underlying cause for aging of humans and other animals. The relationship between mitochondrial dysfunction and ske letal muscle aging has been supported by numerous studies, which found that skeletal muscle aging is accompanied by a decline in mitochondrial density and

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18 function, and dysregulation of mitochondrial quality control processes such as mitochondrial biogenes is, fission and fusion, and mitophagy, the selective autophagic removal and degradation of damaged mitochondria (reviewed by Seo et al. 2010 and Johnson et al. 2013). The goal of this review is to summarize the current knowledge about the age related ske letal muscle dysfunction, with specific focus on mitochondrial dysfunction, and some potential mechanisms underlying the decline in mitochondrial content and function. Aging R elated D ecline in S keletal M uscle M ass and F unction Skeletal muscle is one of th e largest organs in the body, comprising nearly 40 50% of total body mass in non obese mammals. It is an exceptionally plastic tissue that can undergo adaptive changes to meet new challenges imposed on it. For example, muscle mass can either increase or decrease in response to metabolic demands like exercise or following a period of inactiv ity, respectively (Romanello & Sandri, 2015). As we age skeletal muscle undergoes progressive changes, primarily involving loss of muscle mass and strength, known as sa rcopenia ( r eviewed by Buford et al. 2010), which becomes more severe with increasing age and can lead to mobility impairment and frailty ( Cruz Jentoft et al. 2010; Marzetti et al. 2013). In humans, skeletal muscle mass declines as much as 3 10% per deca de after the age of 25 ( Rogers et al. 1990; Short et al. 2005; Johnson et al. 2013), with a dramatic acceleration of decline after the age of 65 (Nair, 2005). Postmortem examination of healthy individuals who died in accidents revealed that both muscle area and fiber numbers started to decrease as early as the fourth decade of life when compared across all ages (Lexell et al. 1988). This seems to be in line with other studies reporting age related decrease in muscle area starting at about 30 y r in healt hy human subjects ranging in age from 18 to 88 y r

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19 (Short et al. 2003 ; Short et al. 2004 ). Moreover, increasing evidence shows the decline in muscle mass is accompanied by a decrease in muscle strength and power. As we a ge, humans lose approximately 1 % of leg lean mass per year and approximately 2.5 4 % in leg strength (Goodpaster et al. 2006). Similarly, Short et al. (Short et al. 2003 ; Short et al. 2004 ) demonstrated a loss of muscle strength (knee extension) concomitant with the decrease in muscle mas s between the ages of 20 and 80 y r This indicates that not only muscle mass declines with age but the muscle quality does as well. However, age related changes vary substantially among individuals likely due to different fitness levels and other environme ntal factors (reviewed by Marzetti et al. 2017). Muscle aging is noted in various species from invertebrates to higher organisms. In the nematode Caenorhabditis elegans aging is accompanied by muscle deterioration and reduced muscle function, which resembled human sarcopenia (Herndon et al. 2002). As C. ele gans age, muscle mass significantly shrinks and sarcomeres progressively disintegrate and become disorganized (Her ndon et al. 2002). A recent study in flies demonstrated similar age related changes in muscle morphology and function (Miller et al. 2008). For example, myofibrils in striated muscle from old flies displayed reduced sarcomere length and increased disorga nization, indicating a loss of sarcomere integrity and acute sarcopenia. Despite vastly different longevity of invertebrate and humans, and essential differences in their muscle fiber types and innervation, similar features of muscle aging have been descri bed. Notwithstanding, only limited data are available for equine skeletal muscle aging. In a study performed on horses of 1 to 21 years of age, the cross sectional area of the longissimus muscle was

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20 et al. 2015). Seemingly in accordance, Betros et al. ( Betros et al. 2002 ) measured lower whole body oxygen consumption (Vo 2 max) and maximum heart rate in aged compared to young horses. It is possible that the decreased aerobic performance capacity of th e older horses was related to decreased muscle mass and function, but many other factors contribute to whole body aerobic performance, and none were investigated in that study. Changes in S keletal M uscle F iber T ype C omposition with A ging Mammalian skeletal muscle is an extremely heterogeneous tissue. It is composed of muscle fibers as well as connective and adipose tissue, and within muscle fibers there are different types that exert specific contractile and metabolic properties. The overall perform ance and function of a muscle are mainly dependent on the individual properties of different fiber types and their proportions within the muscle. Major differences between muscle fiber types are related to the contractile element myosin and the different i soforms of its heavy chain component (MyHC). The different MyHC isoforms are characterized by specific mode s of energy production and muscle fiber function. In large animals, skeletal muscle fibers are classified based on the MyHC isoform expressed. Type I fibers, or slow twitch fibers, are associated with predominantly oxidative energy metabolism; type IIX (or historically IIB) fibers, or fast twitch fibers, are associated with predominantly glycolytic energy metabolism; and type IIA fibers, an intermediat e fiber type, exploit both oxidative and glycolytic energy production. Of note, the number of MyHC isoforms expressed in skeletal mus cle varies between species, and small mammals are reported to express a forth isoform (type IID) (Talmadge and Roy, 1993). In addition to pure fiber types, expressing only one MyHC isoform, hybrid fibers exist, which express more than one isoform. Those are more

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21 prevalent in muscles undergoing transition, such as during aging or a daptation to exercise (Pette & Staron, 2000). F urther, muscle groups very rarely express only one fiber type; instead, they are comprised of a combination of the three fiber types. Aging has been associated with a decline in muscle fiber number, which is though t to be the principle cause of age related loss of muscle mass. A significant decline in the number of muscle fibers was observed in elderly humans. By counting the number of fibers in vastus lateralis muscle of men age d from 30 to 74 y r Lexell and colleagues (Lexell et al. 1986 ; Lexell et al. 198 8 ) reported that the number of muscle fibers in the oldest man was ~25% less compared to the youngest. In humans, the decline in muscle fiber number is more profound in type II fibers. It has previously been reported that type IIX and IIA fiber numbers decline with age (Lexell et al. 1988), whereas the type I fibers are less affected Moreover, the preferential atrophy of type II fibers, that is the loss of fiber s is likely to cause a concomitant increase in percentage of type I fibers. Studies on human skeletal muscle aging suggested that the percentage of type I fibers increases with aging, with ~40% type I fibers in individuals in their twenties, and ~70% type I fibers in individuals in their sixties (Gollnick et al. 1972; Larsson & Karlsson, 1978). In addition to fiber number, muscle fiber size is also affected by age, with 10 40% smaller type II fibers observed in elderly compared to young individuals (Fronte ra et al. 2000). A number of studies investigated the association of muscle fiber atrophy and loss of motor neurons. This idea was first suggested by Gutmann and Hanzlikova ( Gutmann & Hanzlikova, 1966), who reported age associated alterations in the motor endplate morphology at the neuromuscular junction. Since that time, evidence has accumulated

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22 showing that functional denervation occurs during the aging process, including a progressive decline in the number of spinal cord motor neurons, a process conside red irreversible, and of functioning motor units (Roth et al. 2000 a ; Roubenoff, 2001). A motor unit consists of one motor neuron innervating a group of muscle fibers. With age, motor neurons die and cause a denervation of the muscle fibers within the moto r unit, which subsequently leads to muscle fiber atrophy and death. Another factor causing muscle fiber atrophy with age is believed to be the impairment of skeletal muscle protein turnover. Muscle mass is determined by coordinated balance between protein synthesis and degradation. This balance was disrupted in aged humans, driven by an overall decrease in muscle protein synthesis rate, including that of specific contractile and mitochondrial proteins ( Yarasheski et al. 2002; Short et al. 2004). The synthesis of mixed skeletal muscle proteins was described to decline by 4% per decade after the age of 20 (Short et al. 2004). A significantly lower synthesis rate of myofibrillar proteins and MyHC was observed with age ( We l le et al. 1993; H asten et al. 2000), which may be related to age associated reduction in either or both gene transcription and translation of MyHC IIX and MyHC IIA (Balagopal et al. 2001). Furthermore, the exercise stimulated response in myofibrillar and mitochondria l protein synthesis following resistance or endurance training (Moore et al. 2009) was attenuated in older humans (Kumar et al. 2009). The reduction in synthesis of specific contractile proteins is likely to contribute to a decline in locomotor function. Besides loss of muscle fibers, aging skeletal muscle is characterized by a fiber type transition. Skeletal muscle is highly malleable and able to adapt to altered

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23 functional demands and stimuli (Flck, 2006). A common adaptation is a change in muscle meta bolism through modifying fiber type composition. Under certain conditions, fiber type transitions occur between fast and slow fiber types, either from fast to slow or vice versa. In general, increased neuromuscular activity or overload induces fast to slow transitions, whereas reduced neuromuscular activity or unloading causes transitions in the opposite direction (Pette, 2002). Aging is associated with a fast to slow shift of muscle fibers, which was demonstrated by Gannon et al. ( Gannon et al. 2009), who observed a drastic increase in slow myosin light chain in aged rat muscle. Similarly, a proteomic study of human skeletal muscle indicated a slower contracting mode of senescent muscle fibers ( Gelfi et al. 2006 ). Of note, this fast to slow fiber type shi ft is affecting mostly IIX fibers, which can be explained by the age related remodeling of motor units that more likely result in denervation of type II muscle fibers (D'Antona et al. 2003 ; Kostek & Delmonico, 2011), whereas type I fibers are less affected. However, the fast to slow fiber type shift does not increase oxidative capacity in older muscle, which could be due to the reduction in oxidative enzyme activity with age (Boffoli et al. 1994 ; Roo yackers et al. 1996 ). Fiber type distribution in skeletal muscle from untrained horses appears to reflect alterations in muscle function with advancing age (Rivero et al. 1993; Lehnhard et al. 2004; Kim et al. 2005). However, the data are contradictory with some studies reporting an increase in oxidative, slow twitch fibers (Rivero et al. 1993), and others suggesting an age related shift from oxidative to glycolytic fibers ( Lehnhard et al. 2004; Kim et al. 2005). In addition, whether an age related alteration in muscle fiber type composition is associated with changes in muscle oxidative

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24 capacity has not been well investigated in the horse ( Rivero et al. 1993; Kim et al. 2005). Changes in M uscle E nergy M etabolism with A ging Skeletal muscle plays a central role in locomotion and thereby in many activities of daily living, and its action, muscle contraction, depends on the presence of the energy rich molecule adenosine triphosphate (ATP). There are several pathways for generati ng ATP in skeletal muscle, such as 1) hydrolysis of phosphocreatine (PCr) through creatine kinase reaction, 2) anaerobic glycolysis, and 3) citric acid cycle (TCA cycle) and oxidative phosphorylation (OXPHOS). The former pathways are located in the cytosol while the TCA cycle and OXPHOS are located in the mitochondria. Muscle aging is commonly accompanied by alteration in bioenergetics. Recent microarray data indicated a reduction in mRNA level of 55 gene s involved in energy metabolism by more than 2 fold during the aging process, which may have negative implications for the bioenergetic capacity of human skeletal muscle. In general, these 55 genes were associated with glycolysis, glycogen metabolism and mitochondrial function (Lee et al. 1999). However, t he effect of aging on bioenergetic capacity remains controversial, which is probably due to differences in physical activity levels of the individuals assessed. Muscle cells store some ATP that can be used for muscle contraction immediately, but this is us ually only enough to last for a few seconds. During short term, intense activities, after these ATP stores are depleted, the phosphocreatine (PCr) system (also called the ATP creatine phosphate system) can provide fuel for an additional 4 6 seconds The PC r system serves as the quickest way to replenish ATP levels by directly transferring a phosphate group from creatine phosphate to ADP to

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25 form ATP. After cellular uptake, about two thirds of the creatine in skeletal muscle is phosphorylated to PCr by creati ne kinase, with the remaining one third remaining as free creatine. As muscle works, PCr is used to replenish ATP, preventing a depletion of ATP levels. Aging is associated with alteration in the PCr system. Mller et al. (Mller et al. 1980) found that intramuscular PCr levels were approximately 5% lower in elderly (52 79 yr) when compared to younger (18 36 yr) adults, while intramuscular free creatine levels were 5% higher in older adults. At the same time, elderly individuals exhibited significantly re duced total adenine nucleotides. Subsequently, it was found that the activity of the key enzyme catalyzing the phosphorylation of creatine, creatine kinase, in heart tissue was lower in older compared to middle aged adults (Kaczor et al. 2006). Aksenov et al. (Aksenov et al. 1997) concluded that the activity of creatine kinase was reduced by reactive oxygen species (ROS) damage. Given that generation of ROS increases with age (Chabi et al. 2008), formation of phosphocreatine may be decreased in the aged population due to an oxidatively damaged creatine kinase. Glycolysis is the second fastest way to regenerate ATP and acts as the predominant source of energy provision to support explosive exercise lasting from about 30 seconds to 2 min. This metabolic pro cess produces a net sum of 2 ATP molecules per molecule of glucose (Baker et al. 2010) by converting a glucose molecule into 2 molecules of pyruvate, 2 molecules of H 2 O, and 2 molecules of NADH. In subsequent steps, pyruvate is either converted to lactate through the so called lactic acid system, or transported into the mitochondria, where it is further oxidized by in the reactions of the TCA cycle. Lanza et al. (Lanza et al. 2005) reported an age related decline in glycolytic flux in skeletal muscle, whi ch is consistent with reduced glycolytic enzyme contents

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26 ( Gelfi et al. 2006; Capitanio et al. 2009) and/or activities (Larsson et al. 1978; Pastoris et al. 2000) observed in older muscle. For example, the activity of glycolytic enzymes, such as lactate d ehydrogenase and hexokinase are markedly decreased in elderly humans (Pastoris et al. 2000; Kaczor et al. 2006). In contrast, other studies have shown little or no age related changes in activity of enzymes involved in anaerobic ATP generation, such as lactate dehydrogenase, creatine kinase and adenylate kinase (Borges & Essn Gustavsson, 1989; Coggan et al. 1992) Aerobic metabolism is the slowest, but most efficient way to regenerate ATP. This oxygen dependent metabolic pathway is responsible for most of the cellular energy produced. The aerobic system uses blood glucose, glycogen and fat as fuels to resynthesize much more ATP compared to the other pathways. Since this mode of ATP regeneration requires oxygen, and many of the aerobic reactions occur in the using glucose and glycogen as fuels for aerobic ATP production, the carbohydrate is broken down into pyruvate through glycolysis, and the pyruvate enters TCA cycle wi thin the mitochondria matrix. When fat is used as a substrate, triglycerides (body fat) are first hydrolyzed into free fatty acid s and glycerol in a process called lipolysis. Free fatty acids can then be transported into mitochondria and oxidized to genera te acetyl CoA, which enters TCA cycle for further oxidation. The reduction equivalents generated during both the oxidation of fatty acids and carbohydrates are then used to reduce molecules of the mitochondrial electron transport chain. In the process of e lectron transport along a series of enzyme complexes, an electrochemical gradient is generated across the inner mitochondrial membrane, which is the driving force for the ultimate phosphorylation of

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27 ATP. The aerobic oxidation of carbohydrate and fatty acid s necessitate the fast supply of oxygen from blood, particularly during high energy demanding exercise. However, in needs at least in part due to a lower muscle capilla ry density. The number of capillaries in contact with muscle fibers decreases by up to 40% with age (Coggan et al. 1992). Moreover, this study (Coggan et al. 1992) also demonstrated that maximal aerobic capacity (measured as VO 2 max ) is 35% lower in old compared to young adults. The effects of aging on mitochondrial respiration will be discussed in the next part of this chapter. A number of different energy substrates can be utilized for muscle contraction, including intramuscular glycogen and triglycerides, as well as blood borne glucose and fatty acids. The preferred and predominant substrate for energy production used by a muscle cell dep end s on the type of activity or exercise. During moderate activity, such as walking, energy is generated almost entirely through the aerobic pathway with fat being the predominant energy source. As speed of locomotion increases, in the horse this would be from a walk to a trot or a canter, both aerobic and anaerobic systems are recruited, with fat and glycogen being the main energy source. When horses are p er forming a fast gallop, as an example of a strenuous, short term locomotor activity, the energy is ma inly generated through the anaerobic pathway, using glycogen/glucose as energy substrates. On the other hand, long term, endurance exercise relies predominantly on fat as the principle energy source. Limited data have been published regarding the effect of aging on the energetic capacity of equine muscle. Similar to what has been found in older humans, a decline in VO 2 max has been observed in old horses

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28 ( McKeever & Malinowski 1997 ). Moreover, age appears to be accompanied by an increase in muscle glycolyt ic capacity, and a concomitant decrease in muscle oxidative capacity in horses (Kim et al. 2005). Changes in M itochondrial F unction with A ging Mitochondria are unique organelles in that they contain their own genome (mtDNA). This small genome (16.5 KB in human) encodes 2rRNAs, 22 tRNAs, and 13 protein subunits (Smits et al. 2010). The remainder of the mitochondrial proteins (> 90%) are encoded b y the nuclear genome (nDNA) and synthesized in the cytosol before being imported into mitochondria ( Johnson et al. 2007; Pagliarini et al. 2008). Most m ammalian cells contain hundreds to thousands of mitochondria, and each mitochondrion holds 2 10 copies of mtDNA. Mitochondria vary considerably in shape and size, but they all have the same basic double membrane system, consisting of an inner and an outer mitochondrial membrane. Together they create two separate compartments: internal mitochondrial matrix and the intermembrane space. The mitochondrial matrix contains the enzymes that metabolize pyruvate and fatty acids to generate acetyl CoA, as well as enzymes of the TCA cycle, in which acetyl CoA is further oxidized and the energy carriers, or reduction e quivalents, NADH and FADH 2 are produced. At the inner mitochondrial membrane, these electron carriers donate electrons to the components of the electron transport system that generates most of the 2 is coupled with the phosphorylation of ADP, this process is also named OXPHOS Besides this principal bioenergetic function, mitochondria also play crucial roles in cell metabolism, apoptotic cell death and intracellular signaling ( Finkel & Holbrook, 2000 ; Ryan & Hoogenraad, 2007 ). Therefore, proper mitochondrial function is essential for maintenance of cellular homeostasis as

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29 well as general organismal health. Not surprisingly, alterations in mitochondria density and/or function are associated with divers e pathologic states (Wallace, 2005). There is substantial evidence that mitochondrial dysfunction occurs with advancing age in a wide range of species, including human ( Cooper et al. 1992; Joseph et al. 2012 ). The mitochondria theory of aging suggests th at accumulated oxidative damage within the mitochondria causes irreversible damage to mitochondrial macromolecules, and eventually leads to cellular aging. The basis for this theory is the fact that mitochondria are both the producer and the target of ROS The mitochondrial respiratory chain, or electron transport system (ETS), is a major site of ROS production in the cell. Even though the OXPHOS process is efficient, a small percentage of electrons can still prematurely reduce oxygen during normal respiration, resulting in the formation of ROS (Liu et al. 2002). ROS produced within mitochondria re presents almost 90% of the total ROS produced in the cell, and it has therefore been suggested that mitochondria a re prime targets for oxidative damage. Over the years, substantial evidence has emerged to support that aging is accompanied by changes in mitochondrial integrity and function, including an increase in mtDNA mutations and deletions, reduction in expression of mitochondrial proteins, reduced mitochondrial enzyme activities, and low ATP production. In genera l, ROS production is found to increase in aged mitochondria (Chabi et al. 2008). The lack of protective histones, the limited efficiency of mtDNA repair system (Linnane et al. 1989), and close proximity to the ETS make mtDNA extremely vulnerable to oxidative damage. It has been shown that oxidative damage to mtDNA increases in an age dependent manner in skeletal muscle (Kujith et al. 2005). One

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30 study found that mtDNA deletions occurs in up to 70% of mtDNA molecules in the skeletal muscle of old individuals over the age of 80 (Chabi et al. 2005). Perhaps the strongest evidence suggesting a causative link between mtDNA mutations and aging comes fr om studies on the mtDNA mutator mice In those studies, mice with deficiency in the proofreading function of mitochondrial DNA polymerase ( POLG) exhibited an accumulation of point mutations and deletions in their mtDNA (Kujoth et al. 2005; Vermulst et al. 2008) which subsequently led to an accelerated aging phenotype. Of note, oxidative damage to proteins and lipids within mitochondria has also been observed (Beal et al. 2002; Muller et al. 2004; Murphy, 2009; Staunton et al. 2011 ), which undoubtedly will impact mitochondrial function as well Coupled with the increase in mtDNA damage, the mtDNA content decreases with age in skeletal muscle from humans (Welle et al. 2003 a ; Short et al. 2005; Menshikova et al. 2006; Lanza et al. 2008) and rodents (Barazzoni et al. 2000). The decline in mtDNA content is believed to be at least in part due to oxidative damage, since the difference in mtDNA content between young and old groups tends to be greater in more oxidative fibers (Barazzon i et al. 2000). If mtDNA content is a valid indicator of mitochondrial content, the mitochondrial content observed by electron microscopy may exhibit a similar decrease with age. In fact, a study using magnetic resonance spectroscopy has shown that the va stus lateralis muscle of people over 60 years of age showed lower mitochondrial density compared to their younger counterparts (Conley et al. 2000), and this decline in mitochondrial density was further confirmed by assessment of mitochondrial density usi ng electron microscopy (Peterson et al. 2012). Moreover, activity of citrate synthase (CS), a valid biomarker of mitochondrial density ( Larsen et al. 2012 ), has also reported

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31 to decrease in skeletal muscle from human ( Houmard et al. 1998 ) and horses ( Kim et al. 2005 ), which may explain a decline in muscle oxidative capacity. Age related changes in mitochondria oxidative capacity have been well documented in human skeletal muscles ( Houmard et al. 1998 ; Conley et al. 2000; Hunter et al. 2002 ). However, these studies are not in complete concordance, possibly due to differences in muscle types investigated and physical activity levels of the subjects. The majority of reports, although not all (Houmard et al. 1998; Capel et al. 2005), associated aging with a decline in activity of several mitochondrial enzymes such as 3 OH acyl CoA dehydrogenase activity ( 3 HADH), CS, complex I III and cytochrome c oxidase ( COX; Coggan et al. 1992; Houmard et al. 1998; Barazzoni et al. 2000; Short et al. 2005 ). These impairments may in turn cause a decline in mitochondria respiratory capacity Short et al. (Short et a l 2005) reported that the maximal capacity of ATP synthesis in skeletal muscle decreased by ~10% per decade, or 5% when normalizing to mitochon drial protein content. Another underlying cause of the reduction in mitochondrial oxidative capacity could be reduction in mitochondrial mRNA or protein. Recent microarray data revealed 957 genes significantly associated with aging, with mRNA levels of com plex I, III, IV and V significantly reduced in skeletal muscle from aged subjects (Su et al. 2015) In addition, aged human skeletal muscle appeared to exhibit alteration in mRNA levels of some nDNA encoded mitochondrial and TCA proteins, including CS (Barazzoni et al. 2000; Welle et al. 2003 b ; Short et al. 2005 ). At the protein level, the content of complex I, III, IV and V were decreased in aged skeletal muscle from both humans (Short et al. 2003; Lanza et al. 2008) and rodents (Lombardi et al. 2009, Picard et al. 2010) but increased amounts of complex II

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32 (Lombardi et al. 2009; O'Connell & Ohlendieck, 2009; Picard et al. 2010). The latter is a nuclear DNA encoded protein, and its increase concomitant to the decrease of complexes that contain mtDNA encoded subunits has been suggested as a compensatory mechanism. Furthermore, skeletal muscle mitochondrial protein synthesis declined with age (Guillet et al. 2004), which is likely another contributor to age related impairment of mitochondrial oxidative capacity and ATP synthesis (Short et al. 2005). Specific mitochondrial enzymes have also been investigated in horses. Kim e t al. (Kim et al. 2005) investigated the age associated change in activity of CS and 3 HADH, and found that CS decreased while 3 HADH remained unaltered In conclusion, reduction in mitochondrial oxidative capacity in older individuals may be due to lower activity of mitochondrial enzymes as well as lower transcript and protein content. Mitochondrial Q uality C ontrol M echanisms As discussed above, mitochondria are continuously challenged by ROS, which cause damage to mitochondrial constituents and ultimate ly mitochondria dysfunction, which in turn leads to increased level of ROS. However, the cell has an intricate surveillance system to maintain mitochondrial integrity and function, a process termed mitochondrial quality control, which includes chaperones a nd proteases to refold or degrade misfolded mitochondrial proteins; the fission/fusion machinery to repair or facilitate the segregation of damaged areas from the mitochondrial network; and mitochondrial biogenesis and mitophagy to regulate mitochondrial t urnover and regeneration. Antioxidant enzymes residing in the mitochondria can also be considered part of the mitochondria quality control system, as they scavenge ROS to reduce oxidative stress and prevent oxidative damage. Moreover, recent research sugge sts a cross talk between mitochondria and other subcellular compartments like the

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33 endoplasmic reticulum to regulate mitochondrial homeostasis ( Nunnari & Suomalainen, 2012; Fu et al. 2013 ). Of note, the activation and relative contribution of each specific quality control mechanism depends on the overall degree of damage. Mitochondrial P roteases and C haperon e s Mitochondria contain a series of chaperone p roteins and proteases. They are considered to be central to mitochondrial quality control, since they play a crucial role in controlling protein homeostasis. Typically, the major function of chaperones is to recognize non native, partially misfolded proteins, and to facilitate their refolding into the f unctional native state. Mitochondria contain several members of the major chaperones, with the most important mitochondrial chaperones belonging to the heat shock protein families Hsp60 and Hsp70 (Luce et al. 2010). However, if mitochondrial damage is exc essive and refolding by chaperones fails to occur, proteases distributed in all mitochondrial compartments degrade misfolded or oxidized proteins into small fragments and amino acids. For this reason, protease mediated mitochondrial quality control is cons idered to be the first line of defense against mild mitochondrial damage. The two best known soluble proteases functioning in the mitochondrial matrix are LONP1 (mitochondrial matrix peptidase LONP) and ClpXP (Caseinolitic peptidase XP). Other proteases re side in the inner membrane space and control the protein turnover in certain mitochondrial compartments. Fission and F usion Mitochondria are highly dynamic organelles that undergo constant fission and fusion to regulate the expansion and morphology of the mitochondrial network. This dynamic nature serves as a second line of defense against damage (Westermann et al. 2010; Palme r et al. 2011) The fission process segregates damaged parts of the

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34 mitochondria network from the healthy part and targets them for removal and degradation through the quality control process autophagy. Fusion, on the other hand, facilitates the exchange of mitochondrial proteins and mtDNA between healthy mitochondria (Van der Bliek et al. 2013) Thus, quality control via the fission fusion machinery is essential for mitochondrial function. Inhibition of fusion induced the accumulation of mtDNA mutations, eventually triggering loss of the mitochondrial genome (Chen et al. 2010). Imbalance between the fission and fusion process causes alterations in mitochondrial structure and function (S hirendeb et al. 2012). For example, loss of fission ability caused a so called hyper fused mitochondrial network, while excessive fission events resulted in small, round, and fragmented mitochondria (Olichon et al. 2003; Chen et al. 2005). What is more, with age, m itochondria appeared abno rmally enlarged and less numerous with age (Terman et al. 2010), indicating that aging may cause a decrease in mitochondrial fission or an increase in fusion events (Yoon et al. 2006). Autophagy Selective mitochondrial autophagy or mitophagy represents another mitochondrial quality control mechanism. When mitochondrial damage is severe and exceeds the mitochondrial repair capacity, selective autophagic removal of mitochondria is induced (Twig et al. 2008). During mitophagy, damaged or dysfunc tional mitochondria are recognized and enveloped by a double membrane structure called phagophore, which subsequently closes to become an autophagosome. The autophagosome then targets its cargo, the enclosed mitochondria, to a lysosome. Upon fusion of auto phagosome and lysosome (autophagolysosome), the lysosomal enzymes catalyze cargo degradation (Kim et al. 2007). Accumulating evidence shows that a

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35 decline in autophagy with age, and that increased autophagy attenuates aging in lab animals ( Wu et al. 2013 ; Schiavi et al. 2015). PINK/Parkin mediated m itophagy is a well studied mechanism reported to regulate mitochondria integrity and function in various species (Clark et al. 2006; Gautier et al. 2008; Narendra et al. 2008). Damaged mitochondria are flagged with PTEN induced kinase (PINK1), followed by recruitment of Parkin to the mitochondria, which initiates the mitophagic process. The accumulation of PINK1 and Parkin on the surface of damaged mitochondria further triggers ubiquitintion of mitochond rial proteins that act as an elimination signal. Ubiquitinated mitochondria are further recognized by an autophagy adapter protein, Sequestosome 1 (p62/SQSTM1). The protein p62/SQSTM1 is tagged to the phagophore membrane by the autophagy protein MAP LC3 II ( Microtubule associated protein 1A/1B chain 3), and facilitates mitochondrial engulfment by the phagophore Autophagosome formation is executed by a collection of several autophagy related proteins (ATG). To date, 35 ATG genes have been identified in yeas t (Nakatogawa et al. 2009), and the associated proteins work in concert or sequence to facilitate the formation and engulfment of cytoplasmic cargo. So far, mitophagy has been the only mechanism know n to degrade mitochondria, and consequently to turn over the whole mitochondrial genome. Therefore, impairment of mitophagy has been implicated in aging and age & Narendra, 2011; Green et al. 2011). In humans, an age related decline in mitophagy was observed (Cavallini et al. 2007) and associated with an accumulation of damaged mitochondria (Masiero & Sandri, 2010). Given that mitochondria are increasingly damaged during the aging process (reviewed by Shigenaga et al. 1994), decreased mitophagic activity

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36 might further exacerbate dysfunction of the mitochondrial population as a whole in older individuals. In contrast, genetically increasing autophagic capacity appears to reduce aging in lab animals (Rana et al. 2013; Wu et al. 2013; Schiavi et al. 2015). Moreover, autophagy can also be stimulated by a number of ways including physical exercise and caloric restriction. For example, caloric restriction leads to an increase in mitophagy and a decrease in levels of senescence markers in rats (Cui et a l. 2013). In rodents, exercise, which is known to reduce the aging phenotype, has been shown to induce an increase in levels of mitophagy proteins in skeletal and cardiac muscle (He et al. 2012; Ogura et al. 2011). Hence, decreased mitophagy likely con tributes to the decline in mitochondrial quality and function that contributes to the aging phenotype. Mitochondrial B iogenesis As one would expect, controlled mitophagy possibly coordinates with mitochondrial biogenesis to sustain a healthy mitochondria pool (Michel et al. 2012). Mitochondrial biogenesis is a complex process, which requires coordinated synthesis and assembly of thousands of proteins encoded by both the nuclear and mitochondrial genomes ( Scarpulla, 2008). In addition, mtDNA replication must be coordinated to meet the requirements of the mitochondria extension. The biogenesis process is predominantly regulated by one of the most studied regulators, Peroxisome proliferator activated receptor (PGC et al. 1999). The transcriptional coactivator regulates various transcription factors involved in mitochondrial biogenesis in mammalian tissues (Wu et al. 1999), includ ing skeletal muscle (Hood et al. 2006). Overexpression of this gene in skeletal muscle increases mitochondrial content (Garnier et al. 2003) and oxidative capacity (Li n et al. 2002), while PGC deficient mice exhibited a significant decrease in muscle mitochondrial biogenesis (Derbr et al. 2012).

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37 Mitochondrial content in human skeletal muscle declines gradually with advancing age ( Crane et al. 2010 ), and an age associated impairment of mitochondrial biogenesis has been reported in animal models (Fan nin et al. 1999; Sugiyama et al. 1993). Concomitantly the level of PGC compared to young ones (Vi a et al. 2009). Consequently, decreased PGC expression has been suggested an important contributor to impaired mitochondrial biogenesis in old individuals. Mitochondrial biogenesis in muscle can be activated by physical exercise and caloric restriction ( L pez LIuch et al. 2006; Piantadosi & Suliman, 2012; Wenz et al. 2013 ). For example, 5 month of endurance exercise induced mitochondrial biogenesis in mice as indicated by increased expression of PGC 1 mitochondrial Transcription Factor A ) (Safdar et al. 2011). Thus, improving mitochondrial biogenesis may present a potential therapeutic target for age related muscle wasting and diseases. Both in vivo and in vitro studies suggest that caloric restriction induced mitoc hondria biogenesis through activating of PGC 1 L pez LIuch et al. 2006 ) Crosstalk between D ifferent M itochondrial Q uality C ontrol M echanisms M itochondria have different defense mechanisms against increasing damage that safeguard mitochondrial integrity. Dysregulation of any these quality control mechanisms is thought to be implicated in skeletal muscle loss and dysfunction observed with aging. In recent years, mounting evidence shows that mitochondrial quality control mechanisms a re not acting independently from each other, but instead exert considerable crosstalk. For instance, in the fruit fly, Drosophila melanogaster cellular levels of TFAM vary in concert with LONP1 levels, suggesting that LONP1 may regulate mitochondrial biog enesis by selective degradation of TFAM. More specifically,

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38 knockdown of LONP1 in D. melanogaster increased and overexpression of LONP1 reduced TFAM abundance and mtDNA copy number ( Matsushima et al. 2010; Matsushima & Kaguni, 2011 ) Increasing evidence furthermore suggests the existence of crosstalk between the mitochondrial fusion/fission machinery and mitophagy (Tanaka et al. 2010; Westermann, 2010) as malfunctioning mitochondria were segregated from the healthy network by fission, and ultimately eliminated through mitophagy (Youle & Narendra, 2011). Thus, it was hypothesized that fission process is a prerequisite for mitochondria selective degradation. Support for this hypothesis came recently from studies that observed PINK/ Parkin mediated mitophagy was prevented when fission was inhibited (Tanaka et al. 2010). Similarly, excessive fusion has been shown to inhibit mitophagy (Twig & Shirihai, 2011) The ubiquitination of fusion proteins such as Mfn1 and Mfn2 by the ubiquitin ligase Parkin, which induced mitophagy (Gegg et al. 2010; Tanaka et al. 2010) is yet another example of crosstalk between mitochondria dynamics and mitophagy. Mitochondria nuclear crosstalk is also known to play a critical role in regulating mitochondrial quality control Under certain conditions, when the protein degradation system is insufficient to control mitochondrial damage, the mitochondria relay signals to the nucleus, which triggers the expression of mitochondrial chaperones and proteases to improv e protein folding or to remove damaged proteins (Jovaisaite & Auwerx, 2015). Moreover, mitochondria signal the expression of transcription factors essential for mitochondrial biogenesis ( Pellegrino et al. 2013). Interestingly, it appears that mitochondria also communicate with the endoplasmic reticulum (ER), which is supported by evidence that ER specific ubiquitin ligase is able to induce a distinct (Parkin independent) cellular pathway to eliminate damaged

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39 mitochondria (Fu et al. 2013 ). Taken together, different mitochondrial quality control mechanism s communicate and crosstalk with one another to maintain mitochondrial integrity and function. However, if the level of damage exceeds the capacity of these protective mechanisms, or if these protective mech anisms fail to work, damaged mitochondria can induce apoptosis and cell death. Aging and M uscle R egeneration C apacity A gradual loss of muscle mass and function, known as sarcopenia, is the most obvious characteristic of aging. The mechanisms responsibl e for sarcopenia in aged muscle are not completely understood, but the age related reduction in muscle regenerative capacity is proposed to contribute to the development and exacerbation of sarcopenia. Reduced regeneration capacity of skeletal muscle cause s prolonged inflammation ( Shadrach & Wagers, 2011 ), which subsequently lead s to loss of muscle mass observed with aging. The regenerative capacity of skeletal muscle is owed to a population of dedicated muscle stem cells, often referred to as satellite cel ls (Sambasivan et al. 2011). Satellite C ells and M yogene sis Satellite cells were first identified in frog striated muscle in 1961 ( Mauro, 1961 ). They are muscle progenitor cells, which reside between the sarcolemma and the basal lamina of muscle fibers and account for 2 5% of sublaminal nuclei in adult muscles (Schultz et al. 1974; Rudnicki et al. 2008). Due to their distinct anatomic position satellite cells appear as small, wedge shaped cells under the electron microscope. They exhibit a high nuclear to cytoplasmic ratio and few organelles, and condensed nuclear chromatin distinguishes them from myonuclei ( Hawke & Garry, 2001). With cell electroporation labeling and lineage tracking technologies, increasing evidence

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40 suggests that satellite cells originate from a dorsal lateral part of the developing somite, known as dermomyotome (Kassar Duchossoy et al. 2005; Relaix et al. 2005 ). During embryonic muscle development, a subset of myogenic progenitor cells that express specific transcription factors (paired box protein 3 and 7, Pax3 and Pax7) arise from the central dermomyotome and remain as an undifferentiated reserve cell populat ion throughout embryogenesis. In the late fetal stage, these cells migrate to the distinct satellite cell niche (Kassar Duchossoy et al. 2005; Relaix et al. 2005). Satellite cells exist in all vertebrate species, but their density varies between muscles, with that in adult soleus muscle was 2 fold higher than that in tibialis anterior muscle or extensor digitorum longus muscle (Schmalbruch & Hellhammer, 1977; Snow, 1983). Even within the same muscle, more satellite cells are found close to slow type I compared to fast type IIA or IIB myofibers ( Gibson & Schultz, 1982; Okada et al. 1984 ). During the past 5 decades, increasing evidence has gre atly supported the notion that satellite cells play a significant role in muscle repair after injury and perhaps in the maintenance of muscle mass (Bir essi & Rando, 2010). In adult muscle, satellite cells are typically mitotically quiescent; however, upon stimulation they are driven out of their quiescent state, and start to proliferate, differentiate and fuse to injured myofibers or form new fibers. Importantly, a small proportion of activated satellite cells exit the cell cycle and return to the quiescen t state to maintain the satellite cell pool to respond to future muscle injury or growth stimuli. The mechanisms underlying satellite cell activation remains largely unknown. However, the satellite cell microenvironment, also called satellite cell niche, s eems to control this transition from quiescence to activation. Some intracellular signals such as nitric oxide cause an increase in hepatocyte growth factor (HGF), which in turn

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41 triggers activation of satellite cells ( Wozniak & Anderson, 2007). In addition fi broblast growth factors (FGFs) are also known to be ind ispensable for satellite cell activation (Jones et al. 2005 ). Satellite cell myogenic potential is regulated by the expression of Pax3/7 and myogenic regulatory factors (MRFs). Paired box protein Pax 7 is a canonical biomarker characteristic for satellite cells, as it is specifically expressed in satellite cells, in both quiescent and prolif erating stages, while it is abs c ent in differentiated muscle cells. The crucial role of Pax7 in postnatal mai ntenance and self renewal of satellite cells ( Seale et al. 2000; Kuang et al. 2006) was supported by a Pax7 mutant study. Shortly after birth, Pax7 mutant mice suffered a progressive loss of satellite cells, which was mainly due to satellite cell death a nd not exhaustive activation and differentiation; and remnant satellite cells exhibited poor muscle regenerative capacity (Kuang et al. 2006). Pax 3, the paralogue of Pax 7, is also expressed in a subset of quiescent and activated satellite cells ( Relaix et al. 2006 ). Pax3 and Pax 7 play distinct roles in activation of the myogenic differentiation factor MyoD but do not interfere with the expression of the other myogenic regulatory factors ( Relaix et al. 2006 ) The differentiation factor MyoD induces the differentiation of proliferating satellite cells. Interestingly, Myf5 (myogenic factor 5, which is up regulated during satellite cell activation ) can compensate for the loss of MyoD, and mice lacking MyoD exhib it normal muscle m orphology but express about 4 fold higher levels of Myf5 (Rudnicki et al. 1992). However, lacking both Myf5 and MyoD leads to a lack of myoblast expansion (Kassar Duchossoy et al 2004), indicating that the expression of at least these genes is required for myogenesis. As a downstream target of MyoD, myogenin is involved in differentiation and fusion of myoblasts into a myotube ( Megeney and Rudnicki, 1995)

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42 The absence of myogenin expression prevented myoblasts from contributing to postnatal muscle growth. In support of the latter, Knapp et al. ( Knapp et al. 2006) found that low postnatal myogenin expression resulted in reduced body weight in mice. Taken together, postnatal muscle growth and regenerati on depends on satellite cells, whose myogenic capacity is regulated by sequential activation and repression of various myogenic regulatory factors (reviewed by Bentzinger et al. 2012). Aging R elated D ecline in S atellite C ell F unction Although skeletal muscle has a remarkable capacity to regenerate after injury thought out most of life and to grow in response to exercise, abnormal muscle regenerative capacity ( Bockhold et al. 1998; Charg et al. 2002) has been observed in aged muscles. In support, a uto grafting experiments with muscle from old rodents indicated that damaged muscle is less frequently replaced, suggesting that aged muscles also have relatively slow muscle repair and regenerative capacity ( Kaasik et al. 2007; Fell & Williams, 2008 ). I n line with regenerative impairment, a recent study suggested that regeneration of muscle contractile proteins was much slower in the old rats compared to young rats (Kaasik et al. 2007). Given the key role of satellite cells in adult muscle regeneration diminution in their myogenic capacity is considered to contribute directly to the decline in muscle regenerative capacity with age. Indeed, numerous studies have shown the age associated reduction in both the number and myogenic properties of satellite c ells in various species ( Roth et al. 2000 b ; Conboy et al 2003; Chakkalakal et al. 2012; Sousa Victor et al. 2014 ). In general, satellite cells account for about 30% of sublaminal nuclei in neonate mice, but this number decreases to only 2 4% in adult m ice ( Hawke & Garry, 2001). What is more, the progressive decrease in satellite cell number limits

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43 muscle regeneration with aging. For example, Collins et al ( Collins et al. 2007 ) reported that the satellite cell content decreased by 50% in old mice (age 2 y r ) when compared to young animals (age 2 mo). Moreover, loss of satellite cells is associated with muscle fiber atrophy, supported by the observation that in elderly humans the type II muscle fiber atrophy was associated with a fiber type specific decli ne in satellite cell content in type II fibers (Verdijk et al. 2014). In addition to the decline in satellite cell number, aged satellite cells show functional deterioration, including loss of stemness ( Sousa Victor et al. 2014 ) and reduced ability to ac tivate, proliferate, and fuse into myotubes ( Charg et al. 2002; Conboy et al. 2003; Baj et al. 2005 ; Day et al. 2 010 ). However, the mechanisms responsible for either the decline in satell ite cell number and function remain unclear. Accumulation of DNA damage ( Rossi et al. 2007 ) and telomere shortening ( Sharpless & DePinho, 2007; Flores et al. 2010; ) might explain the decline in satellite cell number and regenerative capacity in sarcopenic muscle (Kadi & Ponsot, 2010). In addition to damage to the sat of differentiation factors, the induction of excessive proliferation might deplete the quiescent pool, as it has been reported for the intestinal stem cell pool in flies ( Rera et al. 2011 ). If this is true for sate llite cells, the maintenance of a quiescent satellite cell pool is also crucial for muscle regeneration (Shefer et al. 2006). However, other studies reported no significant decline in the satellite cell pool in aged mice and humans (Roth et al. 2000 b ; Co nboy et al. 2003; Wagers et al. 2005 ), a discrepancy that could be due to experimental differences such as age or gender of the individuals investigated, or different subpopulations of muscle stem cells examined.

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44 The impact of aging on satellite cell myogenic potential is still debated. There is evidence supporting a decrease of both proliferation and differentiation capacity of satellite cells with age ( Schultz & Lipton, 1982; Conboy & Rando, 2005; Conboy et al. 2003; Collins et al. 2007 ; Buford et al. 2010 ), while other research found that proliferative capacity remains constant throughout life (Renault et al. 2000; Hawke & Garry 2001). In the last decade, with the development of molecular and cellular biology techniques, the mechanisms underlyin g age related changes in satellite cell function have been extensively investigated. It has become more clear that the age associated deterioration of satellite cell function arises from both intrinsic changes within the satellite cell itself and influence s of extrinsic factors comprising the satellite cell niche. Satellite cells in aged muscle exhibit decreased responsiveness to repair stimuli. More specifically, they fail to respond adequately to endocrine and paracrine signals that would induce their activation and proliferation in younger muscle (Gopinath & Rando, 2008; Kuang et al. 2008). Recent studies have shown that a key activator of sat ellite cells, Notch ligand Delta like 1, failed to respond to repair stimuli in aged satellite cells (Conboy et al. 2005), but, in contrast, significantly increased in young muscle to regulate satellite cell activation and proliferation ( Conboy et al. 20 02) Lower level of Transforming growth factor beta 1 ( TGF ), an important myogenic regulatory cytokine, has been observed in cells obtained from old individuals compared to that of their young counterparts ( Carlson et al. 2009 b ; Alsharidah et al. 2013 ) In contrast, FGF2 level in the satellite cell niche appeared to be higher in old muscle, which lead to loss of quiescence and self renewal capacity, and eventually resulted in diminished regenerative capacity. Suppression of the FGF2 signaling pathway r escued these

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45 defects (Chakkalakal et al. 2012). The strongest evidence supporting the notion that alterations in the satellite cell microenvironment regulate satellite cell function comes from cross transplantation and parabiotic studies, in which engraft ment of satellite cells obtained from young mice into old mice improved muscle regenerative capacity (Conboy et al. 2005; Villeda et al. 2011 ; Lavasani et al. 2012 ), while transplantation of satellite cells derived from old into young hosts had the oppo site effect ( Carlson & Faulkner, 1989 ). Taken together, these findings suggest that the extracellular environment that defines the satellite cell niche is an important determinant for muscle regeneration through its effect on the satellite cell itself. As noticed in some transplantation studies, the post transplantation regenerative capacity of geriatric satellite cells is reduced compared to that of adult satellite cells when transplanted into animals of the same age (Sousa Victor et al. 2014), suggest ing that in addition to extrinsic satellite cell niche conditions, factors intrinsic to the satellite cell might play a role in age dependent impairment of regenerative capacity. Intrinsic factors such as genomic integrity, mitochondrial dysfunction and ep igenetic changes are thought to contribute to the decline in satellite cell myogenic capacity with aging. Epigenetic remodeling or DNA damage to specific genes, such as key cell cycle regulators, control s cellular fate or turnover of satellite cells, and m ight therefore impair regenerative capacity of various cell types including muscle and its stem cell populations (Lansdorp & Peter, 2007; Sousa Victor et al. 2014). Research on mitochondrial function revealed that satellite cell differentiation relies on functional mitochondria, and a decline in their density and/or function may be a potential signal for satellite cell dysfunction. A marked stimulation of mitochondrial biogenesis accompanies

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46 the differentiation of myoblasts into myotubes both in vivo ( Dugu ez et al. 2002) and in vitro (Moyes et al. 1997; Leary et al. 1998). Myoblasts with either deficiency in respiratory capacity ( Herzberg et al. 1993) or in mitochondrial protein synthesis (Korohoda et al. 1993) fail to differentiate into myotubes. Furthermore, in mice and humans, impaired autophagic activity was observed in aged satellite cells ( Garcia Prat et al. 2016 ) In conclusion, diminished satellite cell function is a consequence of multiple age related impairments, and each of these intrins ic or extrinsic alterations may affect muscle regenerative capacity, and thereby disturb muscle mass maintenance. As the population of old er horse s grows, their owners strive for improved health an d management strategies that will enable continued use of their older horses for athletic and/or recreational activities. Many horses are still actively working even at ages greater than 20 years old ( Brosnahan & Paradis, 2003a; Brosnahan & Paradis, 2003b ). Research involving rodents and human s shows an age rela ted decrease in skeletal muscle aerobic capacity regenerative capacity and mitochondrial functi on as discussed above Yet, few similar studies have been done in the horse. The overall goal of this dissertation was to examine age related changes in muscle energy metabolism, regenerative capacity, and mitochondrial function in equine skeletal muscle, and to explore potential mechanisms that contribute to the age associated alteration in mitochondrial content and quality in equine skeletal muscle.

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47 CHAPTER 2 EFFECTS OF AGING ON MITOCHONDRIAL FUNCTION IN SKELETAL MUSCLE OF AMERICAN QUARTER HORSES Background Aging is a process during which tissue functions progressively decline (Anton et al. 2015; Crescenzo et al. 2015) Age associated impairment of structure and function consequences for the well being of the aging individual. In humans, loss of muscle mass and function, namely sarcopenia, starts to become appare nt in the fourth decade of life, progressively worsening in individuals over the age of 70 ( Buford et al. 2010 ) These age related changes lead to reduced mobility, and consequentially to an impairment of independent living ( Anton et al. 2015 ) While res earch on skeletal muscle aging in humans, primates, and rodent models has led to numerous publications, equine skeletal muscle aging is less explored. The horse is among the most athletic animals (Taylor et al. 1981 ) and used as a companion animal in work, competitive and leisure activities. In recent decades the number of horses over the age of 15 years, which is considered aged, has steadily increased in the US (McKeever et al. 2002 ) in part due to more specializ ed care and veterinary advances. Several recent surveys have pointed out that horses can still actively work and perform athletically well into their 20s ( Brosnahan & Paradis, 2003a; Brosnahan & Paradis, 2003b ) an age physiologically analogous to a 65 yea r old human. The growing population of aged and geriatric horses in the US and their continued use for work and athletic activities raises Reprinted with permissi on from Li C, White SH, Warren LK, Wohlgemuth SE (2016). Ef fects of aging on mitochondrial function in skeletal muscle of American American Quarter Horses. J Appl Physiol (1985) 121 299 311.

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48 the question whether age associated changes in skeletal muscle that have been observed in aged humans and laboratory animals also occur in the horse. Moreover, with the willingness of owners to use and work with this older horse population, understanding the physiology of equine aging is warranted. There are some studies that have documented skeletal muscle functional ca pacity in horses (reviewed by Leisson et al. 2008 ), and most of these studies used horses younger than 12 years of age ( Essn et al. 1980; Lopez Rivero et al. 1991; Roneus et al. 1991; Roneus, 1993; Barrey et al. 1999) therefore not addressing the pr oblems and phenomena associated with aging into the geriatric stage of life. In the present study, we recruited sedentary horses be tween the age of 17 and 25 yr to investigate the impact of physiological aging on skeletal muscle function in comparison to a group of young horses. Skeletal muscle is highly malleable and able to adapt to altered functional demands and stimuli (Fl ck, 2006 ) A common adaptation is a change in muscle fiber metabolism through modifying fiber type composition. Skeletal muscle myosin heavy chain an essential component of the contractile apparatus, exists in different isoforms, which are associated with mode of energy production and muscle fiber function. In large animals, based on the MyHC isoform present, skeletal muscle fiber s are classified as type I, associated with predominantly oxidative energy metabolism; type IIX (or historically IIB), associated with predominantly glycolytic energy metabolism; or type IIA, which is an intermediate fiber type, exploiting predominantly ox idative energy production with functional features characteristic of glycolytic fibers. Importantly, hybrid fibers exist, which express more than one isoform, and are more prevalent in muscles undergoing transition, such as aging, or exercise adaptation (r eviewed by Pette & Staron, 2000 ).

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49 Further, muscle groups very rarely express only one fiber type; they are comprised of a combination of the three fiber types. Fiber type distribution in skeletal muscle has been investigated in untrained horses to reflect alterations in muscle function with advancing age. However, the data are contradictory, with some studies reporting an increase in oxidative fibers (Rivero et al. 1993 ) and others suggesting an age related shift from oxidative to glycolytic fibers (Lehnh ard et al. 2004; Kim et al. 2005) In addition, whether an age related alteration in fiber type composition is associated with changes in muscle oxidative capacity has not been well investigated in the horse ( Rivero et al. 1993; Kim et al. 2005 ) Skeletal muscle oxidative metabolism occurs in the mitochondria. Mitochondria are prominent in skeletal muscle and provide ATP for muscle function through the process of OXPHOS carried out by the complexes ETS Increasing evidence shows that aging is associated with compromised capacity for OXPHOS in human muscle ( Cooper et al. 1992; Joseph et al. 2012 ) most likely due to a decline in mitochondrial density and/or function (Short et al. 2005 ) Numerous mitoc hondrial components have been tested during the past two decades as biomarkers of muscle oxidative capacity in human ( Mogensen et al. 2006; Ritov et al. 2006; Boushel et al. 2007 ) as well as in horses, such as CS 3 HADH ( Kim et al. 2005; Revold et al. 2010 ) COX (White et al. 2015 ) and succinate dehydrogenase (Serrano et al. 2000 ) Among them, activity of CS, COX, and 3 HADH have been used as biomarkers of mitochondrial content and oxidative capacity, respectively ( Coggan et al. 1992; Rimbert et al 2004; Larsen et al. 2012 ) Moreover, OXPHOS is a complicated process, which requires the coordinated interaction of multiple enzyme complexes. The mere assessment of enzymatic function

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50 of individual components of this system may be insufficient to reve al how well t he system functions as a whole, and whether and where age related defects occur ( Saks et al. 1998; Villani et al. 1998; Kunz, 2003 ) High resolution respirometry (HRR) with saponin permeabilized muscle fibers allows the study of OXPHOS in sk eletal muscle mitochondria in situ Oxygen consumption of mitochondria still situated within the muscle fiber can be sensitively monitored and evaluated by titrating energy substrates, ADP, complex inhibitors, and uncouplers to the permeabilized fibers. To date only a few groups have appli ed this technology in horse studies to investigate the effect of training on muscle function ( Votion et al. 2010; Votion et al. 2012; White et al. 2015 ) We are unaware of any studies using HRR with permeabilized muscle fibers to study the impact of agi ng on integrated mitochondrial respiratory function in horse skeletal muscle. The objective of this study was to examine the age associated changes in skeletal muscle fiber type composition and mitochondrial function in the Quarter Horse. We sampled the G LU and TRI muscles of the extremities with distinct locomotor functions attributed to and associated with different muscle fiber type composition (van den Hoven et al. 1985 ) To address the question whether alterations in fiber type composition, determin ed electrophoretically, are associated with changes in muscle oxidative capacity, both the classical spectrophotometric method of enzyme activity assays and the more comprehensive HRR were used to assess mitochondrial OXPHOS capacity. Furthermore, the comp arison of mitochondrial function between GLU and TRI muscle allowed us to evaluate muscle specific responses to aging.

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51 Materials and Methods Animals All procedures performed in this study were approved by the University of Florida Institute of Food and Agricultural Sciences Animal Research Committee. A total of 34 Quarter Horses including 24 young (1.8 0.1 years old, 14 fillies and 10 geldings) and 1 0 aged subjects (17 25 years old, 9 mares and 1 gelding) were utilized in this study. Horse body condition score (BSC) was assessed using the Henneke scoring system with a scale from 1 to 9 (1 = emaciated and 9 = extremely obese) (Henneke et al. 1983 ) Co mpared to the young horses, which had a BCS of 5.00, the aged horses had a significantly higher average BCS (5.80 0.25; P < 0.001). All horses used in this study were owned by the University of Florida. None of the young horses had undergone any type of conditioning; and none of the aged horses had been athletes in the past, nor been used in competitions, and had been exposed to only low intensity exercise/training in the past, if any. None of the horses used in this study had received forced exercise 6 m onths prior to the study. Skeletal M uscle S ampling Skeletal muscle samples were taken from the right or left GLU (n = 34: 24 young + 10 aged) and from the TRI of a subset of young horses (n = 12) and of all aged horses (n = 10) under local anesthesia using a 14 gauge SuperCore TM b iopsy needle (Angiotech, Gainesville, FL, USA). The GLU muscle was located on the croup by tracing approximately one third down a line from the tuber coxae to the tailhead; and the long head of the TRI muscle was located at th e intersection of a line traced between the scapulohumeral and radiohumeral joint and a vertical line extending down from the tricipital crest of the scapula. Briefly, the sample collection site (3 3 cm) was shaved,

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52 and the skin was cleaned with 7.5% pov idone iodine and rinsed with 70% ethanol. Horses were sedated with approximately 0.3 mL of detomidine hydrochloride (Dormosedan, Pfizer Animal Health, Exton, PA) administered i.v. The sampling site was then anesthesized by subcutaneous injection of 0.2 0. 3 mL 2% mepivacaine hydrochloride (Carbocaine V, Pfizer Animal Health, Exton, PA). The muscle sample (approx. 30 to 40 mg tissue) was taken at a standardized depth of 5 cm within the respective muscle through an initial skin puncture created with a 14 gau ge needle and subsequent insertion of the biopsy needle. In order to collect a sufficient quantity of tissue sample, the biopsy needle was reinserted 2 3 times into the same initial puncture at different angles (same depth). Samples were immediately transf erred into 1 mL ice cold biopsy preservation solution (BIOPS; 2.77 mM CaK 2 EGTA, 7.23 mM K 2 EGTA, 5.77 mM Na 2 ATP, 6.56 mM MgCl 2 2 O, 20 mM Taurine, 15 mM Na 2 Phosphocreatine, 20 mM Imidazole, 0.5 mM DTT, and 50 mM MES; pH 7.1 (Saks et al. 1998 ) ) for subsequent respiration measurement on fresh tissue sample. The remaining tissue sample was snap frozen in liquid nitrogen and stored in a dry shipper (MVE SC4/2V, CHART, Inc., Ball Ground, GA, USA) for transport to the laboratory and later stored at 80 C Preparation of P ermeabilized M uscle F ibers The preparation of permeabilized muscle fibers was described previously (Kuznetsov et al. 2008 ) Briefly, muscle samples were immersed in ice cold BIOPS solution during transportation to the laboratory. Before further preparation of fibers, fat and connective tissue were removed, and muscle fibers gently separated using two pairs of forceps. Myofiber bundles were transferred to 1 mL fresh BIOPS solution eabilized for 30 min at 4

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53 C on a rotator. After the permeabilization, the myofibers were rinsed in ice cold mitochondrial respiration medium (MiR05; 110 mM sucrose, 60 mM potassium lactobionate, 0.5 mM EGTA, 3 mM MgCl 2 2 O, 20 mM taurine, 10 mM KH 2 PO 4 2 0 mM HEPES, and 1 g/L BSA; pH 7.1) on a rotator for 10 min at 4 C. Mitochondrial respiration of these permeabilized myofibers was immediately assessed using HRR. High R esolution R espirometry Mitochondrial respiration (O 2 flux) was determined in duplicate using the high resolution respirometer Oroboros O2k (OROBOROS INSTRUMENTS, Innsbruck, Austria) at 37C and in hyperoxic conditions (220 2 ), according to methods previously described (Kuznetsov et al. 2008 ) Unles s stated otherwise, the concentrations of the following reagents used are final concentrations in the respirometer chamber. Permeabilized fibers (2 3 mg wet weight) were added to the respirometer chamber containing 2 mL of MiR05. The respiration medium was supplemented with 20mM creatine to saturate mitochondrial creatine kinase, which facilitates mitochondrial ADP transport ( Saks et al. 1991; Walsh et al. 2001 ) Oxygen flux was determined with the following titration protocol (Fig ure 2 1): electron flow through complex I (CI) of the ETS ( OXPHOS capacity C I P C I ) was supported by the NADH linked substrates glutamate (10 mM) and malate (2 mM) (LEAK respiration, L), followed by addition of adenosine diphosphate (ADP, 2.5 mM) to stimulate respiration (OXPHOS CI P CI ); the addition of succinate (10 mM) supported convergent electron flow through complex I and II (CII) of the ETS (OXPHOS capacity C I+II P C I+II ); additional ADP (1.25 mM) and succinate (5 mM) were then titrated to evaluate whether OXPHOS capacity could be increased any further. No further increase was observed, indicating that OXPHOS capacity was measured at a saturating ADP concentration. Subsequent

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54 a ddition of cytochrome c (cyt c membrane. O nly samples with response to cyt c below a 15% increase in respiration were included in the analysis. Titration of the uncoupler carbonyl cyanide 4 respiration (ETS capacity; E). The coupling control ratio L/P was calculated as the ratio of LEAK to OXPHOS capacity (L/P C I+II ) and the P C I+II /E coupling control ratio (P/E) was calculated as the ratio of OXPHOS capacity (P C I+II ) to ETS capacity. Sample P reparation and SDS PAGE for M yosin H eavy C hain A nalysis Frozen muscle was powdered in liquid nitrogen using a BioPulverizer (BioSpec Products, Inc., Bartlesville, OK, USA). Tissue powder was subsequently homogenized with 30 volumes of extraction buffer (50 mM Tris HCl, pH 8.3, 10 mM EDTA) using a glass tissue grinder (Kontes Dual, size 20, Kimble Chase, Vineland, NJ, USA). The muscle homogenate was transferred to a microcentrifuge tube and diluted with equal volume of protein denaturation buffer (4% SDS, 20% glycerol, and 125 mM Tris HCl, pH 6.8). The sample was then heated at 50 C for 20 min and centrifuged at 14,000 x g for 20 min (101) Protein content was determined using the Bicinchoninic Acid (BCA) Inc., Rockford, IL, USA). Samples were stored in equal volume of glycerol at 20 C until further processing. MyHC isoforms in muscle homogenates were analyzed using a slightly modified protocol described previously ( Talmadge & Roy, 1993 ) Briefly, the protocol was modified as follows: 1) samples were heated at 95 C for 3 min; 2) electrophoresis (SDS electrophoretic protein separation, the gel was stained with Coomassie Bri lliant blue

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55 R250 (Bio Rad Laboratories, Inc., Hercules, CA, USA) for 2 h and the proteins visualized using a cooled CCD camera and acquisition software (G BOX imaging system, Syngene, Frederick, MD, USA). To verify the nature of the separated bands, a sepa rate gel electrophoresis with a representative subset of muscle sample homogenates was performed. Instead of Coomassie staining, the proteins were transferred to a PVDF membrane using a semi dry blotter at 20 V for 60 min (Trans Blot Bio Rad Laboratories, Inc., Hercules, CA, USA). The nature of the protein bands were confirmed as MyHC IIA, MyHC IIX and MyHC I isoforms by immunoblotting using primary myosin heavy chain antibodies SC 71 (MyHC IIA, 1: 150), 6H1 (MyHC IIX, 1: 100) and BA D5 (MyHC I, 1:100) (al l from Developmental Studies Hybridoma Bank ), and HRP conjugated goat anti mouse IgG secondary antibody (1: 80,000, Sigma, St. LABO RATORIES, Inc., Burlingame, CA, USA) and the chemiluminescence captured with the G Box imaging system. The intensity of the Coomassie blue stained protein bands (Fig ure 2 2) as well as the chemiluminescence intensity was analyzed and quantified by densitom Syngene). For each sample, percentages of each MyHC isoforms were calculated as a percentage of the total (100%) MyHC isoforms in each lane. Spectrophotometric D etermination of E nzyme A ctivities Enzymatic activities of CS, COX, and 3 HADH in muscle homogenates were measured as previously described ( Fong & Schulz, 1978; Spinazzi et al. 2012 ) using a microplate reader (Synergy HT, BioTek Instruments, Winooski, VT, USA). Briefly, CS activity was as sessed at 412 nm by measuring the initial rate of reaction of free CoA SH with DTNB; COX activity was determined by measuring the maximal, linear rate of

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56 oxidation of fully reduced cytochrome c at 550 nm; and 3 HADH was assayed by measuring the oxidation o f NADH at 340 nm using acetoacetyl CoA as substrate. Statistical A nalyses D ata are reported as mean s SE with the number of samples per group noted in the figure legends. Statistical analyses were performed using SigmaPlot version 12.0 (Systat Software, Inc, San Jose, CA). All data were compared between different age groups and across skeletal muscles using two way analysis of variance (ANOVA) followed by Student Newman Keuls multiple comparison tests. For data that did not express a normal distribution (L/P), a log transformation was performed prior to statistical analysis, but the original data were presen ted in the graph. In all comparisons, differences with P reported if 0.05 < P < 0.1. Results Aging and M uscle F iber T ype C omposition The composition of MyHC isoforms was significantly alte red in muscles from aged compared to young horses, with increase of MyHC I and IIA, and decrease of MyHC IIX, ind ependent of muscle type (Table 2 1 ; main effect of age; MyHC I: P = 0.002, MyHC IIA: P = 0.032, MyHC IIX: P = 0.004). More specifically, in GLU muscle the percentage of MyHC I was significantly higher in aged compared to young horses ( P < 0.001). In addition, there was a trend for a decreased percentage of MyHC IIX in aged GLU muscle ( P = 0.092), while the proportion of MyHC IIA was unaltered ( P = 0.520). In TRI muscle, no difference in percent MyHC I was detected between the two age groups ( P = 0.224). However, aged horses exhibited a higher percentage of MyHC IIA ( P = 0.023), and a lower percentage of MyHC IIX ( P = 0.012) relative to young anima ls.

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57 When GLU and TRI muscles were compared, the MyHC composition significantly differed between muscles, and this difference was maintained in both age groups (main effect of muscle type, P < 0.001 for all fiber types). More specifically, GLU muscle had a lower percentage of MyHC I and MyHC IIA, and a higher percentage of MyHC IIX compared to TRI Effect of A ge on M itochondrial D ensity and E nzyme A ctivity We found a main effect of age on CS activity per mg tissue ( P = 0.01 4; Figure 2 3A). Specifically, CS activity did not differ between age groups in GLU ( P = 0.230), but was lower in TRI muscle from aged horses ( P = 0.023). When the two muscles were compared, CS activity in the TRI muscle was approximately 2 fold greater compared to the GLU (main effect of muscle type: P < 0.001), independent of age, which is consistent with the almost 2 fold difference in percentage of presumably mitochondria rich (type I and IIA) muscle fibers observed between the two muscles. There was a main effect of age on COX activit y per mg muscle tissue ( P < 0.001; Fig ure 2 3B). Specifically, COX activity per mg tissue decreased significantly with age in both muscles ( P < 0.001). Furthermore, COX activity per mg tissue was significantly lower in GLU compared to TRI muscle (main effe ct of muscle type: P < 0.001). When COX activity was normalized to CS activity to reflect the oxidative capacity on the mitochondrial level (Figure 2 3C), there was a significant effect of age (main effect of age: P < 0.001). S pecifically, COX activity per mitochondria l unit was 47% lower in aged GLU ( P < 0.001), and 30% lower in aged TRI muscle ( P = 0.003). When muscles we re compared, COX activity per mitochondrial unit was lower in TRI compared to GLU muscle (main effect of muscle type: P < 0.001).

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58 Activity of 3 HADH per mg muscle tissue was not affected by age in either GLU or TRI muscle (main effect of age: P = 0.125; Figure 2 3D). Compared to the GLU the TRI muscle exhibited higher 3 HADH activity per mg muscle tissue (main effect of muscle type: P < 0.001). When 3 HADH activity was evaluated per Mt unit by normalizing i ts activity to CS activity (Figure 2 3E), 3 HADH activity was significantly elevated with age in both muscles (main effect of age: P < 0.001; GLU : P = 0.023; TRI : P < 0.001). When muscles were compared, 3 HADH activity per mitochondrial unit was lower in TRI compared to GLU muscle (main effect of muscle type: P = 0.044). Effects of A ge on M itochondrial R espiration There were no apparent age associated changes in mass specific mitochondrial respiration (O 2 flux; pmol O 2 /s/mg wet weight) in either GLU or TRI muscle for any of the assessed respiratory states (LEAK: GLU : P = 0.487, TRI : P = 0.606; P CI : GLU : P = 0.820, TRI : P = 0 318; P CI+II : GLU : P = 0 386, TRI : P = 0 422; ETS capacity: GLU : P = 0 738, TRI : P = 0 918; Figure 2 4A D). When muscles were compared, significant differences were detected between GLU and TRI muscle (Figure 2 4A D). Specifically, the TRI exhibited higher OXPHOS capacity (P CI and P C I+II ), and ETS capacity when compared to the GLU muscle (main effect of muscle type: P < 0.001 for all). When O 2 flux was normalized to CS activity (Figure 2 4E H), we found a significant effect of age for all b ut OXPHOS capacity supported by glutamate, malate and succinate (P CI+II ), independent of muscle type (main effect of age: LEAK: P = 0.038; P CI : P = 0.045; P CI+II : P = 0.073; ETS capacity: P = 0.007). In particular, P CI and ETS capacity tended to be higher in aged GLU muscle ( P = 0.086 and P = 0.085, respectively; Figure 2 4F, H), and ETS capacity was significantly higher in aged TRI muscle ( P = 0.033; Figure 2 4H) when compared to the young counterparts. When the

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59 two muscles were compared, OXPHOS capacity s upported by glutamate and malate (P CI ) ( P = 0.025), glutamate, malate and succinate (P CI+II ) ( P = 0.027), and ETS capacity ( P = 0.038) were significantly higher in GLU compared to TRI muscle ( P values reflect m ain effects of muscle type; Figure 2 4E H). These findings are consistent with COX activity per mitochondrial unit, and suggest that the higher oxidative capacity of TRI muscle results from a higher mitochondrial density rather than from oxidative capacity of individual mitochondria. Effect of A ging on C oupling C ontrol R atios We used the O 2 flux in different respiratory states to calculate coupling control ratios. The L/P coupling control ratio (LEAK/P CI+II ), an indicator for coupling of oxygen consumption and phosphorylation, was between 0.06 and 0.07 for young TRI and young GLU muscles, and between 0.08 and 0.11 for aged TRI and GLU muscles, suggesting a tight coupling of mitochondr ia in permeabilized fibers (Figure 2 5A). There was a main effect of age on the L/P coupling control ratio ( P = 0.016). Compared to young horses, the L/P ratio was significantly increased by 49 % in GLU ( P = 0.019), but remained unchanged in TRI muscle ( P = 0.263). When muscles were compared, the L/P ratio did not differ between GLU and TRI muscle independent of age (main effect of muscle type: P = 0.326). To examine the extent to which oxidative phosphorylation exploits the full capacity of the ETS, we calculated the P/E coupling control ratio (P CI+II /fully non coupled ETS capacity; Figure 2 5B). We found that P/E was significantly affected by age (main effect of age: P = 0.011), but when the effect in the individual muscles was evaluated, there was only a tendency for the P/E ratio to decrease with age (GLU : P = 0.076; TRI :

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60 P = 0.057). No dif ference in P/E was detected between the two muscles (main effect of muscle type: P = 0.854). Discussion In the present study, we compared skeletal muscle metabolic phenotype by analyzing muscle fiber type composition and mitochondrial OXPHOS capacity in Quarter Horses of two different age groups. For this comparison, we chose the GLU and the TRI muscle based on their functions. The GLU muscle is located in the hind limb and used for explosive propulsive movement, while the TRI muscle is a postural muscle located in the forelimb, where it supports the body weight during long periods of standing. These different functional demands dictate a different muscle fiber type composition and energy metabolism. Based on differential susceptibility of muscle fiber typ es to age related changes that has been reported in humans and rodent models ( Houmard et al. 1998; Phillips & Leeuwenburgh, 2005; Joseph et al. 2012 ) we expected to see different responses to age in fiber type composition and mitochondrial function in t hese two types of equine muscles. MyHC isoform appears to be an appropriate marker for muscle fiber type classification, and has been used in numerous studies to characterize skeletal muscle fiber type composition ( Rivero et al. 1997; Andersen, 2003; Jose ph et al. 2012 ) Our findings in the Quarter Horse are in agreement with previous findings in horses (van den Hoven et al. 1985 ) Both GLU and TRI muscles had a relatively high content of MyHC II isoforms (88 96%), which might partially explain the great capacity for explosive speed of this horse breed. However, GLU muscle had relatively more MyHC IIX fibers than the TRI muscle while the TRI muscle contained a greater percentage of MyHC I fibers. This is consistent with observations in other horse breeds at the same biopsy sampling

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61 depth (van den Hoven et al. 1985 ) functional demands for explosive power for propulsion (GLU ) or for postural support (TRI ), respectively. In our study, aging was associated w ith a shift in skeletal muscle fiber type composition toward a MyHC I or IIA phenotype, but with a differential response in the two muscles investigated. Aging has been shown to affect fiber type distribution in various horse breeds ( Essn et al. 1980; Rivero et al. 1993; Lehnhard et al. 2004; Kim et al. 2005 ) ( Table 2 2 ). For example, Rivero et al and Kim et al ( Rivero et al. 1993; Kim et al. 2005 ) found a decline in percentage of type I fibers with age, but no correlation of age and percentage of type IIA or IIX fibers in the Semitendinosus muscle from a group of horses of different breeds, with ages ranging from 2 to over 30 years. Likewise, Lehnh ard et al. (Lehnhard et al. 2004 ) reported a decrease in MyHC I and IIA, and an increase in MyHC IIX content in GLU from old (20+ years old) compared to young (4 8 years old) Standardbred mares. On the contrary, our results suggest an age associated increase in MyHC I fibers in the GLU and MyHC IIA fibers in the TRI muscle, respectively, concomitant with a relative decrease in MyHC IIX fibers in both muscles. This is in agreement with findings in humans (Welle et al. 2000 ) rodents ( Sugiura et al. 1 992; Sullivan et al. 1995; Pehme et al. 2004 ) and two reports in horses ( Essn et al. 1980; Rivero et al. 1993 ) ( Table 2 2 ). Indirect support for a fiber type shift to a more oxidative phenotype comes from a study in aging humans by Lanza et al. (Lanza et al. 2005 ) Using phosphorous magnetic resonance spectroscopy of contracting skeletal muscle in vivo the authors demonstrated that older men relied more on oxidative phosphorylation and less on glycolysis for ATP production compared to young men The

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62 discrepancies between our results and some previously published findings in equine muscle could have resulted from the choice of breed and the age range investigated. Compared to other breeds, the skeletal muscle of the Quarter Horse is rich in type IIX fibers (Valberg, 2013 ) which could have accentuated the shift in fiber type distribution compared to breeds with a more balanced distribution. In addition, the age of the young horses investigated in this study was about 2 years, an age at which the skeletal muscle is still de veloping (Rietbroek et al. 2007 ) which could have affected the outcome of our measurement in comparison to other studies. In rodents, fiber type distribution of different muscles was differentially affected by aging. For example, in rats age related alt erations were only observed in slow twitch Soleus muscle but not in fast twitch Tibialis anterior muscle of the same individual (Larsson & Edstrom, 1986 ) A study in Standardbred trotters ( Essn et al. 1980 ) assessed muscle spec ific fiber type adaptions to the aging process and reported that GLU and TRI muscles had the greatest increase in oxidative fiber types among studied muscles. There are several potential mechanisms that could underlie the age associated change in relative fiber type comp osition. A selective age related susceptibility to atrophy of a specific fiber type could lead to relative increase s in the other fiber types ( Ciciliot et al. 2013 ) Selective atrophy of mainly type II(X) fibers and predominantly fast twitch muscles has been widely accepted and described to occur in rodents ( Thompson & Brown, 1999; Lowe et al. 2001 ) and humans (Joseph et al. 2015 ) For example, in elderly humans, t he elevated relative increase of type I fibers was mainly caused by a selective atrophy of type II fibers (Larsson et al. 1978 ) Muscle fiber

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63 atrophy observed with age can, at least in part, be driven by several mechanisms, such as a decrease in protein s ynthesis (reviewed by Yarasheski, 2003 ), or mitochondria dependent and independent apoptosis ( Marzetti et al. 2010 ) To the best of our knowledge apoptosis or protein degradative mechanisms in skeletal muscle have not yet been investigated in the aged ho rse. Wagner et al. (Wagner et al. 2013 ) reported that whole body protein synthesis was unaffected by age in mixed breed horses, but that there seems to be a disturbance of skeletal muscle specific protein synthesis (namely in the mTOR signaling pathway). However, muscle structure, muscle fiber features and other effectors of the mTOR pathway such as atrophy and autophagy proteins were not assessed in this study. A comprehensive assessment of aging equine skeletal muscle will be useful for a deeper understa nding of the underlying causes of age related muscle weakness, especially when development of interventions to attenuate age related decline in physical performance is desired. An alternative explanation for the altered fiber type distribution is an age i nduced fast to slow fiber type transition within a given fiber (reviewed by Larsson & Ansved, 1995 ). This transition could be caused by denervation of type II fibers followed by either atrophy of the denervated fiber or by subsequent reconstitution of larger motor units with slow motor neurons, inducing a fiber type switch ( Pette & Staron, 2000 ) Studi es in rodents undergoing fast to slow conversion indicated that the transition in MyHC isoforms follows a sequential order from MyHC IIB to MyHC IIX to MyHC IIA to MyHC I ( Jaschinski et al. 1998 ) Based on our data, we speculate that if there was an age r elated conversion of fiber types in the muscle of Quarter Horses, it occurred to a differential degree or in a differential time line in the two muscles. With MyHC IIX

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64 decreasing in both muscles, the GLU displayed an increase in MyHC I and the TRI in MyHC IIA portion, which suggest that GLU could have been affected at an earlier time point or to a more severe degree compared to TRI Similar muscle specific changes associated with age were suggested by Essn and coworkers ( Essn et al. 1980 ) in a study on racing horses, in which the authors compared age related responses in fiber type distribution in the GLU muscle with that in other muscles (includin g TRI muscle ). In this study, the GLU muscle was the only muscle that displayed an increased type I/II ratio with age. Further studies need to be conducted to inves tigate a differential time line or severity of changes in fiber type distribution between dif ferent muscles. In addition, in order to distinguish between loss of fibers and fiber type c onversion, total fiber number, cross sectional area of fibers, and muscle weight would be ideal parameters to determine. However, the lack of practicality prohibits some of those measures in the study of live animals in which only small muscle biopsies can be acquired. In summary, our results are in agreement with extensive literature on humans and rodents. However, at this point, we cannot distinguish between selective fiber atrophy or fiber type conversion, nor the development of hybrid fibers. In additi on, it needs to be emphasized that the literature is all but consistent in the observation of age related changes in fiber type composition. Purves Smith et al. ( purves Smith et al. 2014 ) recently critically reviewed and evaluated the methodology leading to the various and often contradictory findings. The authors concluded that the presence of hybrid fibers, which is rarely assessed, may draw a different picture of age related changes in fiber type contribution altogether, and that the lack of identification of hybrid fibers could account for the contradictory results on the effect of age on fiber type composition.

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65 Our next aim was to compare the observed shift to a higher percentage of t ype I or type IIA fibers with the metabolic phenotype on a subcellular level. Specifically, we asked whether the age induced increase in type I or type IIA fibers was concomitant with altered function and OXPHOS capacity of skeletal muscle mitochondria; an d whether muscle oxidative capacity was altered through modifying mitochondrial content, mitochondrial respiratory capacity, or both. C itrate synthase activity was used as a representative biomarker of mitochondrial density and total cristae area ( Larsen et al. 2012 ) When we compared the two muscles, we found that CS activity was 2 fold higher in TRI muscle compared to GLU muscle, which corresponds with the 2 fold higher proportion of fibers with a presumably predominantly oxidative metabolism (MyHC IIA + MyHC I) in TRI muscle compared to GLU muscle. We furthermore found that GLU and TRI muscles were affected differently by age. More specifically, CS activity decreased with age in the TRI muscle but not in the GLU m uscle. A decline in CS activity has not often been documented in old sedentary horses (Kim et al. 2005 ) But findings similar to ours were reported in human studies that revealed an age associated decline in CS activity in the oxidative portion of the Gas trocnemius muscle (Houmard et al. 1998 ) but not in the relatively more glycolytic Vastus lateralis ( Grimby et al. 1982; Houmard et al. 1998 ) The explanation for this divergent change in CS activity between different muscles with aging is not clearly e vident, but it may be partially explained by MyHC distribution. In the Quarter Horse, CS activity was unaffected by age in GLU muscle. However, this mitochondria rich type I fibers, suggesting that the increased proportion of type I fibers

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66 did not cause a proportional increase in mitochondrial de nsity. What is more, CS activity diminished with age in the TRI muscle despite the observed relative increase in MyHC IIA content. In contrast to our findings, Kim et al. (Kim et al. 2005 ) reported that CS activity in equine Semimembranosus a predominant ly glycolytic muscle, was negatively correlated with age, and concomitant with a decreased MyHC I proportion. However, we this mitochondrial density, was proportional to th e decline in oxidative fibers. The underlying cause for the age associated decline in CS activity ( Hebert et al. 2015 ) and mitochondrial density described for human muscle ( Conley et al. 2000 ) might be a decreased rate of mitochondrial protein synthesis ( Guillet et al. 2004 ) or alterations in mitochondrial biogenesis, which is associated with mitochondrial protein synthesis (Joseph et al. 2012 ) Alternatively, an age associated imbalance between autophagic removal of mitochondria, namely mitophagy, and mitochondrial biogenesis would diminish mitochondrial density. However, impaired rather than increased mitophagy has been widely observed in aged muscle (reviewed in Hepple, 2014 ). To the best of our knowledge neither mitochondrial biogenesis nor mitophag y have been assessed in investigations. Next we addressed the question of whether the age related decrease in mitochondrial density (deduced from CS activity) correlated with de creased oxidative capacity. We assessed muscle mitochondrial function, namely OXPHOS capacity, by both spectrophotometric measurement of COX activity in muscle homogenates, and HRR of skeletal muscle mitochondria in situ In addition to OXPHOS capacity, we also

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67 assessed activity of 3 oxidation pathway to evaluate the capacity to metabolize fatty acids for energy production. There is some debate on whether differences in muscle tissue OXPHOS are due to mitochondrial content and/or co mposition of muscle fiber types, or function of the individual mitochondria, or both. Initially, it was suggested that differences in muscle respiration may be attributed solely to the differences in mitochondrial quantity ( Hoppeler et al. 1987; Schwerzma nn et al. 1989 ) Recently, a greater number of studies have revealed that differences in muscle oxidative capacity depended on the function of the mitochondria themselves ( Jackman & Willis, 1996; Amara et al. 2007 ) Here, we present COX and 3 HADH activities and O 2 flux relative to muscle weight, as well as per unit mitochondria. The normalization of mitochondrial functional markers to a mitochondrial marker such as CS activity has been recommended by Pesta and Gnaiger ( Pesta & Gnaiger, 2012 ) in ord er to separate the effects of mitochondrial quality (function) from mitochondrial quantity. In agreement with studies in humans ( Rimbert et al. 2004 ) and horses (Kim et al. 2005 ) our data on 3 HADH activity suggest that fatty acid oxidation capacity wa s not impaired on a muscle level, and even elevated on a mitochondria level in muscles from our aged Quarter Horses. However, others reported an age related d ecline of 3 HADH activity in humans ( Coggan et al. 1992 ) and a decreased ability to utilize fatty acids for mitochondrial respiration in mice (Johnson et al. 2015 ) The preserved ability to metabolize fatty acids suggested by our data in aged horses seems to be concurrent with the structural shift towards a h igher percentage of type I and IIA muscle fibers. Gueugneau et al. (Gueugneau et al. 2015 ) reported a higher lipid content in type I and IIA compared to type IIX fibers in human Vastus lateralis muscle, and an increase in

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68 intramyocellular lipid content wi th age, although this was not accompanied with a fiber type shift. Cytochrome c oxidase activity has been validated as a marker of mitochondrial function, specifically of OXPHOS capacity ( Larsen et al. 2012 ) Our data showed that COX activity declined with age in both the GL U and the TRI muscle, whether normalized to wet weight or to mitochondrial unit, suggesting an impairment of OXPHOS capacity in older horses not only through decline in mitochondrial number but also through impaired function of the m itochondria themselves. Joseph et al. (Joseph et al. 2012 ) found that COX activity (per unit muscle weight) in elderly, low functioning humans was lower than in that of young individuals, but the authors did not measure any indicator of mitochondrial density. We observed an over 40% reduction in COX activity with age in both muscles. This decline was more severe in the GLU especially when the COX activity was normalized to mitochondrial unit. Given that GLU mitochondrial density was not severely affected by age, it appears that the oxidative capacity is primarily impaired on the individual mitochondrial level. On the other hand, in the TRI muscle, which displayed a significant decrease in mitochondrial density, the impairment of COX activity on the mitochondrial level seemed less severe. In the present investigati on, we did not measure production of ROS or markers of (oxidative) damage to mitochondrial components, which could potentially underlie the pronounced reduction in COX activity in GLU muscle. Compared to oxidative muscles, muscles of a more glycolytic phen otype have been shown to produce more ROS, to have lower free radical scavenging capacity, and to consequently display higher oxidative damage such as lipid peroxidation ( Anderson & Neufer, 2006; Picard et al. 2008; Picard et al.

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69 2011 a ) Comparing the two muscles, independent of age, an interesting finding was that TRI muscle exhibited higher COX activity than GLU muscle when normalized to muscle weight, but lower COX activity per mitochondrial unit, which suggests that in TRI muscle the higher COX acti vity on the tissue level was achieved by a higher mitochondrial content. To further explore the muscle specific impairment of mitochondrial function with aging we measured mitochondrial respiratory function in situ using HRR. Although our results demonst rated that COX activity in skeletal muscle homogenates was dependent on age and muscle type, we cannot deduce whether this alteration reflects a change of muscle mitochondrial OXPHOS. Saponin permeabilized muscle fibers allow examination of the integrative mitochondrial function in a relatively preserved cytoarchitecture ( Kuznetsov et al. 1998; Saks et al. 1998 ) Our mass specific respirometry data demonstrated no age related difference in muscle respiratory capacity across all respiratory states, indicat ing that the capacity for muscle OXPHOS remained high in both muscles from our aged Quarter Horses. When expressed relative to CS activity, LEAK, OXPHOS (P CI ) and ETS capacity were elevated with age (significant main effect of age), independent of muscle type, although only aged TRI muscle displayed a significant increase in ETS capacity. Products of oxidative stress are known to accumulate with age, and are reported to activate mitochondrial uncoupling proteins (UCPs) and the adenine nucleotide transporte r (ANT) ( Echtay et al. 2002; Echtay et al. 2005 ) Both could have caused an increased proton leak, and hypothesis of aging suggests that mild uncoupling, particularly in ag ed tissue, protects against mitochondrial ROS production and subsequent oxidative damage to

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70 mitochondria and other cellular components ( Brand, 2000; Greco et al. 2003; Speakman et al. 2004 ) The elevated LEAK respiration in aged muscle explains only a mi nor part of the concurrently increased ETS capacity. It is possible that this increase reflects on a functional level the increasing reliance on oxidative energy production in aged fibers. Jacobs et al. ( Jacobs et al. 2013 a ) recently showed in mice that mitochondria specific ETS capacity was elevated with age in the primarily glycolytic Gastrocnemius muscle, but not in the primarily oxidative Soleus muscle. Both mass specific and mitochondria specific activated respiration s upported by complex I and II substrates (OXPHOS capacity, P CI+II ) were not affected by age, which is contrary to the age related decline in COX activity that we determined in muscle homogenates. A similar case is reported by Chabi et al. who assessed state 3 respiration (OXPHOS in the presence of glutamate, malate and ADP) of isolated mitochondria and COX activity in muscle homogenates ( Chabi et al. 2008 ) The control of mitochondrial respiration is shared between all of the complexes and electron carriers in the respiratory electron transport system ( Rossignol et al. 2000 ) suggesting no rate limiting step per se In addition, it was evident in human mitochondria that maximal complex IV activity is in excess of what is required for OXPHOS ( Gnaiger et al. 1998; Villani et al. 1998 ) This is supported by the observation that inhibition of complex IV activity had to exceed a critical value (40 60%) to cause a detectable decrease in mitochondrial respiration in both isolated mitochondria ( Villani & Attardi, et al. 2001 ) and permeabilized muscle fibers (Kunz et al. 2000 ) These suggestions could explain why we did not detect a decrease in OXPHOS with aging in either muscle as a whole when we assessed mitochondrial respiration with substrates supporting

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71 complex I and II. We did not assess mitochondrial respiration supported by fatty acid oxidation. In rodents, fatty acid supported respiration was impaired with age ( Hey Mogensen et al. 2012; Johnson et al. 2015 ) while fat oxidation capacity was similar in muscle from young and old sedentary men ( Rimbert et al. 2004 ) The unaltered 3 HADH activity on a whole muscle level observed here suggests that the capacity to oxidize fatty acids was preserved in skeletal muscle form aged horses. Future s tudies oxidation is correlated with utilization of this energy by the Mt electron transport chain. The difference between mass and mitochondria specific respiration when comparing GLU and TRI muscle s is consistent with our results for COX activity. Triceps brachii muscle had higher mass specific respiratory rates across all states (except LEAK), which could be due to its considerably higher mitochondrial density. However, mitochondria specific respiration in TRI muscle was lower (with the exception of LEAK). This difference in mass and mitochondria specific respiration between different equine muscles is similar to observations in humans and laboratory animals ( Jackman & Willis, 1996; Amara et al. 2007 ) For example, in old mice mass specific activated respiration was higher and mitochondria specific activated respiration lower in the oxidative Soleus compared to the glycolytic Gastrocnemius muscle ( Jacobs et al. 2013b ) which suggests a mitoc hondrial specialization with muscle and/or muscle fiber type. Moreover, the finding that mitochondria in glycolytic muscles possessed a higher oxidative capacity than those in oxidative muscles is consistent with observations in other mammals ( Jackman et a l. 1996; Picard et al. 2008; Jacobs et al. 2013a ) Specific mitochondrial phenotypes have been postulated to exist across skeletal muscle types,

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72 with different composition and morphology. For example, others have reported that mitochondrial composition can vary by differential expression of respiratory complex subunit isoforms to sustain the tissue specific energy demands ( Grossman & Lomax, 1997; Huttemann et al. 2001; Kunz, 2003 ) ; and Jacobs et al. (Jacobs et al. 2013a ) determined differential protein expression of all ETS complexes in mouse skeletal muscle homogenates of three different muscles. We thus speculate that mitochondria from GLU muscle might express more COX protein, reflected in the higher mitochondria speci fic COX activity and respiratory capacity compared to TRI mitochondria. At this point, we cannot substantiate our speculation, and further analyses will have to quantify mitochondrial components in the different muscles. In addition, mitochondria vary morp hometrically between slow and fast fibers. Fast twitch muscle mitochondria are arranged in a thinner and longer reticular network, while mitochondria in slow twitch muscle possess a thicker and more truncated network (Ogata & Yamasaki, 1997 ) which could b e related to the lower mitochondria specific respiratory capacity in TRI muscle. We found no difference in L/P or P/E ratio between the two muscles. A study with elite athletes pointed out that differences in mitochondrial OXPHOS capacity and mitochondrial coupling control were apparent only when substrates for both complex I and II are provided ( Jacobs & Lundby, 2013 ) We hence calculated L/P and P/E using OXPHOS capacity with a physiological substrate combination that supports electron flow through CI and CII (P CI+II ). In the young horses, both muscles exhibited low L/P ratios, indicating a good coupling between respiration and phosphorylation, and the L/P ratios for young horse in our study are similar to those reported by Votion et al. ( Votion

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73 et al. 2012 ) for mature horses with the same respiratory substrates (note: these authors determined the respiratory control ratio (RCR); and 1/RCR=L/P). Although there was a main effect of age on L/P, this control ratio increased significantly only in GLU but not in TRI muscle, further confirming that mitochondria from the GLU muscle are more susceptible to age related damage, which is consistent with the COX activity data. Moreover, a compromised coupling of respiration and phosphorylation is in agreement with th e generally elevated LEAK respiration that we observed with age (independent of muscle type). Age affected the P/E phosphorylation control ratio independent of muscle type (main effect of age). In young horses, P/E was around 0.75 in both muscles, consiste nt with a report on healthy, untrained mature horses with a P/E ratio of 0.8 (Votion et al. 2012 ) The tendency for an increasing limitation of OXPHOS capacity by the phosphorylation system, indicated by reduced P/E, together with the elevated ETS capacit y observed in aged horses, suggests that even though aged horses displayed an elevated maximal ETS capacity, constraints of the phosphorylation system may underlie a lower relative activated respiration, thereby possibly limiting the energy supply in the a ging muscle. However, a limited mitochondrial ATP generation might be necessary to reduce the production of ROS (Brand, 2000 ) ATP production relies on a high proton gradient, which may, in turn, be associated with ROS generation. In this situation, an ele vated proton leak, suggested by the higher LEAK respiration in the older muscles, might help to limit oxidative stress and damage. The data on mitochondrial respiration on the tissue as well as mitochondrial level suggest that despite significant decline in mitochondrial content (CS activity) and COX activity as well as shift in MyHC isoform composition, integrated mitochondrial funct ion

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74 and intrinsic mitochondrial functionality were not negatively impacted in skeletal muscle from Quarter Horses about 20 years of age. Several interpretations come to mind: 1) In this study, we assessed integrated mitochondrial function in situ in permea bilized muscle fibers, a technique that helps to attenuate or even prevent disruptions due to preparative technique (Picard et al. 2011 b ) But similar to assessment of isolated mitochondria, the mitochondria in situ were deprived of their intra and extra cellular milieu, which could have affected mitochondrial function in vivo (reviewed in Hepple, 2014 ). 2) It is possible that the Quarter Horse, which can remain active well beyond the age assessed in this study, does not respond to aging with impaired oxid ative capacity like it has been described for rodents and humans. And lastly, 3) the horses investigated in our study might not have reached the age at which integrated and/or intrinsic mitochondrial function becomes overtly impaired, and were instead at a transition age, displaying cellular adaptions rather than impairments, similar to observations made in aging rhesus monkeys (Pugh et al. 2013 ) Taken together, future studies are needed to test those possibilities, for example by using in vivo non invas ive technologies to assess oxidative capacity within the feasibility of application in large animals, and by expanding the age range of subjects.

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75 Table 2 1. Fiber type composition of g luteus medius and t riceps brachii from American Quarter Horses MyHC Isoforms (%) g luteus medius t riceps brachii Main effect of muscle type, P value Young (n =24) Aged (n =9) Young (n =11) Aged (n =9) MyHC I 3.39 0.35 6.68 0.87*** 10.57 0.99 11.85 0.71 P < 0.001 MyHC IIA 21.35 0.85 23.18 2.65 33.79 1.22 41.42 4.53* P < 0.001 MyHC IIX 75.26 0.92 70.14 2.82 55.64 1.59 46.73 4.92* P < 0.001 Values are means SE P < 0.1, *** P < 0.001 (Young vs. Aged within GLU); P < 0.05 (Young vs. Aged within TRI).

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76 Table 2 2. Effect of age on fiber type composition in equine skeletal muscle Horse breed Age Muscle Methodology Effect of age Ref. SB 2 mo to 28 yr GLU, TRI Histochemistry GLU: type I, IIA type IIB Essn et al. 1980 AL, A 2 3 yr vs. 10 24 yr GLU Histochemistry %type I, IIA %type IIB Rivero et al. 1997 SB 4 8 yr vs. 20 + yr GLU S DS PAGE & Coomassie blue staining %type I, %type IIX Lehnhard et al ., 2004 TB, SB, QH, CB 2 yr to 30 yr S S DS PAGE & Coomassie blue staining %type I Kim et al. 2005 Horse breeds are as follows: TB, Thoroughbred; SB, Standardbred; QH, American Quarter Horse; CB, crossbred; AL, Andalusian; A, Arabian. Muscles investigated are as follows: GLU, gluteus medius ; TRI, triceps brachii ; S, semimembranosus Arrows indicate incr eased, decreased, or unchanged percentage of fiber type.

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77 Figure 2 1 Respirometric protocol with permeabilized fibers from American Quarter Horse gluteus medius muscle. Shown is a typical trace of oxygen consumption after permeabilized fiber preparation with glutamate/malate and succinate substrate combinations to support electron flow through complex I (CI) and complex II (CII), respectively, of the mitochondrial electro n transport system (ETS), and its activation by ADP. Cytochrome c was added as a quality control (see text for details), FCCP to induce uncoupling and evaluate ETS capacity, and antimycin A (inhibitor of complex III of the ETS) to evaluate residual oxygen consumption (ROX). The blue line represents the oxygen concentration (nmol/ml), the red line represents the muscle mass specific O 2 flux (pmol O 2 s 1 mg wet wt 1 ; negative slope of the blue line normalized to tissue weight). Marked sections correspond to s teady state fluxes at different coupling states (L, P, and E; see text for explanations).

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78 Figure 2 2 Representative SDS polyacrylamide gel stained with Coomassie blue following electrophoretic separation. Bands shown are MyHC isoforms typ e I, type IIA, and type IIX, in GLU muscle from young (Young GLU) and aged (Aged GLU), and TRI muscle from young (Young TRI) and aged (Aged TRI) American Quarter Horses. The middle lane shows the molecular mass markers at 2 50 and 150 kDa.

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79 Figure 2 3 Enzyme act ivities in muscle tissue homog enates from American Quarter Horses. A ) activity of CS per milligram tissue of GLU and TRI from young ( n = 24 for GLU n = 11 for TRI ) and aged ( n = 9 for GLU n = 10 for TRI ) horses. B E ) activity of COX ( B and C ) and 3 HADH ( D and E ) normalized to milligram tissue ( B and D ) and to CS activity ( C and E ) in GLU and TRI from young ( n = 23 24 for GLU n = 11 12 for TRI ) and aged ( n = 9 10 for GLU n= 9 10 for TRI ) horses. Values are means SE. Open bars repre sent young horses; solid bars, aged horses. Young vs. aged: P < 0.05, ** P < 0.01, *** P < 0.001 GLU vs. TRI: § P < 0.05, §§§ P < 0.001.

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80 Figure 2 4 Mitochondrial respiration of permeabilized skeletal muscle fibers from American Quarter Horses Mass specific O 2 flux (A D; pmol O 2 s 1 mg 1 wwt), and O 2 flux normalized to CS activity (E H; pmol s 1 U 1 CS), respectively, with LEAK respiration (A and E), OXPHOS capacity CI (B and F), OXPHOS capacity CI+II (C and G), and maximal ETS capacity (D and H) are shown Values are mean s SE; n = 17 18 (Young GLU), 12(Young TRI), 9 (Aged GLU) and 9 (Aged TRI). Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.1, P < 0.05. GLU vs. TRI: P < 0.1, § P < 0.05, §§§ P < 0.001.

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81 Figure 2 4. Continued

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82 Figure 2 5 Mitochondrial coupling control ratios of permeabilized skeletal muscle fibers from American Quarter Horses A) L/P coupling control ratio (LEAK/P CI+II ) of GLU and TRI from young (n = 18 for GLU, n = 12 for TRI) and aged (n = 8 for GLU, n = 9 for TRI) horses B) P/E coupling control ratio (P CI+II /ETS capacity) of GLU and TRI from young (n = 18 for GLU, n = 12 for TRI) and aged (n = 9 for GLU, n = 9 for TRI) horses Values are mean s SE. Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.1.

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83 CHAPTER 3 SKELETAL MUSCLE FROM AGED AMERICAN QUARTER HORSES SHOWS IMPAIRMENTS IN MITOCHONDRIAL BIOGENESIS AND AUTOPHAGY Background Mitochondria serve as the combustion engines of life, providing the ATP necessary for skeletal muscle contraction. They do so through OXPHOS in which ATP and CO 2 are produced at the expense of nutrient substrates and molecular O 2 Given healthy mitochondrial population is critical to ensure efficient energy supply. In human and rodent models, extensive investigation has revealed changes in mitochondrial content and function in a variety of disease models. For example, data collected in old individuals suggest that r educed skeletal muscle mitochondrial content and quality is associated with aging ( Short et al. 2005; Picard et al. 2010; Hepple, 2014 ). The t otal mitochondrial content of a cell is tightly regulated by two opposing cellular processes, mitochondrial biogenesis and mitochondria selective autophagy ( mito phagy) ( Mi shra & Chan, 2016; Wai & Langer, 2016 ). Mitochondrial biogenesis, the creation of new mitochondria, is responsible for replenishment of new functional mitochondria. On the other hand, mitophagy, one of the key mitochondrial quality control mechanisms, sequ esters and degrades dysfunctional or aged mitochondria to maintain a healthy mitochondrial population. Thus, the balance between mitochondrial renewal and elimination of damaged mitochondria is essential for maintaining a certain level of healthy mitochond ria to meet energy demands (revi ewed by Palikaras et al. 2015 ). It is not surprising that an impaired balance between these opposing processes can result in numerous pathological conditions, including aging, in diverse organisms ranging from yeast to mamm als ( Preston et al. 2008; Artal Sanz & Tavernarakis, 2009 ;

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84 Kaeherlein, 2010 ). Mitochondrial biogenesis is a complex process, which requires coordinated synthesis and assembly of thousands of proteins encoded by both the nuclear and mitochondrial genomes ( Scarpulla, 2008 ). In addition, mitochondrial DNA (mtDNA) replication must be coordinated to meet the requirements of the new mitochondrial generation. Growing evidence demonstrates that mitochondrial content in human skeletal muscle declines gradually with advancing age ( Crane et al. 2010 ). Concomitantly, an age associated impairment of mitochondrial biogenesis capacity has been reported in animal models ( Sugiyama et al. 1993; Fannin et al. 1999 ). Mitophagy is a specific form of autophagy that selective ly degrades dysfunctional mitochondria. The mitochondria are sequestered by a unique double membrane organelle, called autophagosome, and targeted to be degraded in the lysosomes ( Ding & Yin, 2012 ). Accumulating evidence shows that a decline in autophagy i s associated with age, and that increased autophagy delays aging in lab animals ( Wu et al. 2013; Schiavi et al. 2015 ). In humans, an age related decline in mitophagy was observed ( Cavallini et al. 2007 ) and associated with an accumulation of damaged mitochondria ( Masiero & Sandri, 2010 ). Given that mitochondria are increasingly damaged during the aging process (reviewe d by Shigenaga et al. 1994 ), decreased mitophagic activity might further exacerbate dy sfunction of the mitochondrial population as a whole in older individuals. Hence, decreased mitophagy likely contributes to the decline in mitochondrial quality and function that contributes to the aging phenotype. Previously we have shown that mitochond rial content and function decreased gradually with advanced age in skeletal muscle from American Quarter Horses (Li et al. 2016 ; Chapter 2 of this dissertation ). However, it is not completely understood what

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85 regulates mitochondrial content and function in the equine. One underlying cause for the age associated decrement of mitochondrial density and function could be an impaired balance between mitochondrial biogenesis and autophagy in muscle from aged horses. Therefore, the purpose of the current study was to explore potential mechanisms that contribute to the age associated decline in mitochondrial content and quality in equine skeletal muscle by examining the factors involved in mitochondrial biogenesis and autophagy. Because of their distinct locomotor f unctions and metabolic properties, we compared GLU and TRI muscles from young and aged American Quarter Horses ( Li et al. 2016 Chapter 2 of this dissertation ). We hypothesized that different muscles respond differently to aging, and that the distinct response is due to differences in the activation of mitochondrial content control mechanisms. A better understanding of the cellular and molecular mechanisms resp onsible for the maintenance of a healthy mitochondrial population in equine skeletal muscle is a prerequisite to design interventions to prolong health and performance of aging horses. Materials and Methods Animals Healthy young (1.8 0.1 years old, 14 fillies and 10 geldings) and aged (17 25 years old, 11 mares and 1 gelding) American Quarter Horses owned by the University of Florida were enrolled in this study. None of the horses had received forced exercise 6 months prior to the study. The Henneke bo dy condition score (BCS) system (Henneke et al. 2014 ) with a scale ranging from 1 (emaciated) to 9 (extremely obese) was used to estimate the horse body condition score of the horses enrolled in the study. The BCS was 5.00 for the young group and 5.80 0.25 for the aged group ( P < 0.001),

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86 respectively. This study was approved by the University of Florida Institute of Food and Agricultural Sciences Animal Research Committee. Muscle T issue S ampling Skeletal muscle microbiopsies were obtained from the GLU (young: n = 2 4; aged: n = 12) and TRI of a subset of young horses (n = 12) and of all aged horses (n = 12) under local anesthesia following the procedure described previously (Li et al. 2016 ; Chapter 2 of this dissertation ) Muscle samples were collected at a sampling depth of 5 cm, using a 14 gauge SuperCore TM Biopsy needle (Angiotech, Gainesville, FL, USA), immediately snap frozen in liquid nitrogen and transported to the laboratory in a dry shipper (MVE SC4/2V, CHART, I nc., Ball Ground, GA, USA), where samples were transferred to a 80 C freezer. Analysis of M tDNA C opy N umber To evaluate mitochondrial content in horse skeletal muscle, the relative amount of mtDNA to nuclear DNA (nDNA) was determined using a CFX Connect real time PCR detection system (Bio Rad, Laboratories, Inc., Hercules, CA). Total DNA was extracted from muscle samples using Wizard Genomic DNA purification kit (Promega pair specific for the mtDNA (NADH dehydrogenase 1, ND1 ) and another specific for the actin, ACTB ), were designed using PrimerQuest (Integrated DNA Technologies, Coralville, IA). The primer sequences for the ND1 GGA TGG GCC TCA AAC TCA A GGA GGA CTG AGA GTA GGA TGA T ACTB CTC CAT TCT GGC CTC ATT GT AGA AGC ATT TGC GGT GGA (reverse). Another primer pair designed within the COX1 ( cytochrome c oxidase subunit 1 ) region

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87 of the mitochondrial genome was also tested to confirm the result assessed with ND1 .The primer sequences used for COX1 CAG ACC GTA ACC TGA ACA CTA C GGG TGT CCG AAG AAT CAG AAT AG The relative amount of ND1 COX1 and ACTB was determined for each sample from a standard curve prepared from a serial dilution of a pool of all the samples. Relative mtDNA copy number was calculated from the ratio ND1 / ACTB and COX1 / ACTB RNA I solation Total RNA was isolated from ~30 mg of snap frozen muscle using RNeasy Plus With the DNA elimination solution included in the k it, genomic DNA was remove d from the samples. Immediately following extraction, the RNA concentration and purity were determined using a UV spectrophotometer (Synergy HT, Bio Tech Instruments, Winooski, VT) by measuring the absorbance at 260 (OD260) and 280 (OD280) nm. All measurem e nts were performed in duplicate Analysis of mRNA E xpression First strand cDNA was synthesized with random primers using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems Inc., Foster City, CA) according to ns. Quantitative real time PCR (qRT PCR) analysis was performed using the CFX Connect real reaction volume containing cDNA, primers, and iTaq TM Universal SYBR Green Supermix (Bio Rad). All samples were analyzed in du plicate simultaneously with a negative control that contained no cDNA. The data were normalized to GAPDH mRNA in each reaction, and results were expressed as a fold change in mRNA compared with expression in GLU muscle from the young horse group. Forward a nd reverse primer

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88 sequences are listed in Table 3 1. Melting point dissociation curves generated by the instrument were used to confirm the specificity of the amplified product. The relative quantification was done using the relative standard curve method. Analysis of P rotein E xpression by Western B lot Frozen muscle samples were cryopulverized using a BioPulverizer (BioSpec Products, Inc., Bartlesville, OK, USA) prior to protein extraction. The cryopulverized tissues were immersed 1:50 (w/v) in lysis buffer (25 mM Tris HCl, pH 7.4, 150 mM NaCl, 5 mM EDTA 1% Triton X 100, 0.1% SDS, 10% glycerol, 1 mM DTT, 0.5% sodium deoxycholic acid ( Wu et al. 2009 ) ) supplemented with 1% Halt TM protease phosphatase inhibitor cocktail (Fisher Scientific, Pittsburgh, PA). After 10 bouts of 1s long sonications (Sonic Disme mbrator, model F60; Fisher Scientific), the tissue lysate was incubated for 2 h on a rotator at 4 C, followed by centrifugation at 12,000 x g for 20 min at 4 C. The supernatant was collected and stored at 80C until further analysis. Protein concentrati on was determined with the Thermo Scientific TM Pierce TM BCA protein assay kit (Fisher Scientific). Samples were diluted with equal volume of Laemmli buffer (Biorad) and heated for 10 min at 95 C. Thirty micrograms protein was loaded and resolved by Sodium dodecyl sulfate polyacrylamide gel electrophoresis using precast gels (BioRad), transferred onto a Polyvinylidene fluoride membrane (PVDF; EMD Millipore, Fisher Scientific), and immunoblotted as previously described ( Wohlgemuth et al. 2010 ). Primary anti bodies used for immunoblotting recognized p62/SQSTM1 (1:300, Sigma Aldrich, St. Louis, MO), Atg5 (1:500, Cell Signaling Technology, Danvers, MA), CS (1:100, Santa Cruz, Dallas, TX), LC3 (1:500, Fisher Scientific), and Atg7 (1:1000, Cell Signaling), and sec ondary antibodies were either horseradish peroxidase or alkaline phosphatase conjugated (Sigma Aldrich). Finally,

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89 DuoLux chemiluminescent/fluorescent substrate for horseradish peroxidase or alkaline phosphatase (Vector Laboratories, Burlingame, CA) was ap plied, and the chemiluminescent signal captured with the digital imager G:Box Chemi XR5 (Syngene, analysis software (Gene Tools, Syngene). Protein level was expressed re lative to total protein loaded, as determined by Ponceau staining. Statistical A nalysis Statistical analysis was performed using SigmaPlot 13.0 software (Systat Software, Inc, San Jose, CA). The normal distribution of the data was examined using the Shapiro Wilk test. For data that did not express normal distribution, the log 10 transformation was successfully used to normalize the data. Data were analyzed using two way analysis of variance, with factors of age ( Young, Aged) and muscle type (GLU, TRI ). Significant main effects and interactions were further tested using Holm Sidak multiple comparison tests. Data a re presented as the mean s SE with the number of samples per group noted in the figure legends. A P statistically significant. Results Mitochondrial C ontent W as D ecreased in A ged S keletal M uscle We have previously reported that activity of CS an enzyme of the TCA cycle located in the mitochondria, declined with age in TRI but no t GLU muscle from American Quarter Horses (Li et al. 2016 ; Chapter 2 of this dissertation ). We show here that, consistent with enzyme activity, the T RI muscle exhibited higher CS protein content than the G LU muscle (main effect of age: P = 0.007). Furthermore, compare d to

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90 young muscle relative CS protein content was lower in aged TRI mus cle ( P = 0.048), but not in aged GLU muscle ( P = 0.469; Figure 3 1). Mitochondrial B iogenesis W as I mpaired with A ge To elucidate possible mechanisms underlying the age related decline in TRI mitochondrial content suggested by both CS activity and content, we examined mtDNA copy number, which has been associate d with mitochondrial content (Larsen et al. 2012 ) and biogen esis ( Clay et al. 2009 ). We determined the content of the mitochondrial DNA encoded gene ND1 coding for NADH: ubiquinone oxidoreductase core subunit 1, as a surrogate for mtDNA copy number, and found that, consistent with CS activity and protein expression, mitochondrial DNA content, as indicated by ND1 gene expression (Figure 3 2A), was ~25% l o wer in aged TRI muscle ( P = 0.040) but not in aged GLU muscle ( P = 0.395) when compared to the respective muscles in young horses, suggesting an age related impairment of mi tochondrial biogenesis in the TRI muscle. In addition, mitochondrial DNA content in TRI muscle was h igher than that of GLU muscle independent of age (main effect of muscle: P = 0.011). We confirmed these results by quantifying content of another mitochondrial DNA encoded gene, COX1 (Figure 3 2B), coding for cytochrome c oxidase subunit 1 (young TRI vs. age d TRI : P = 0.017; young GLU vs. aged GLU : P = 0.305; main effect of muscle: P = 0.020). In addition, CS protein content and mtDNA ( COX1 / ACTB ) correlated significantly (R 2 = 0.3632; P = 0.020; Figure 3 2C) in TRI muscle, with low citrate synthase concurring with low mtDNA copy number. To further test if any key factors that regulate mitochondrial biogenesis might be altered in aged skeletal muscle, mRNA expression of Peroxisome proliferator activated

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91 receptor gam ma coactivator 1 ( PGC Figure 3 2D), Nuclear Respiratory Factor 1 ( NRF1 ; Figure 3 2E) and mitochondrial Transcription Factor A ( TFAM ; Figure 3 2F) was measured. Gene expression of a PGC a master regulator of mitochondrial biogenesis in mammals ( Ru as et al. 2012 ), was down regulated in aged TRI muscle (~ 60%, P = 0.022), but unchanged in the GLU ( P = 0.407 ). Concordantly, the transcript level of TFAM acting downstream of PGC Wu et al. 1999 ) was decr eased in aged TRI muscle ( P = 0.037), but unchanged in aged GLU muscle ( P = 0.534). Nuclear Respiratory Factor 1 (also acting downstream of PGC et al. 1999 )) mRNA was not alt ered by age in either muscle (TRI : P = 0.376; GLU : P = 0.513). When GLU and T RI muscles were compared, the mRNA expr ession of PGC NRF1 and TFAM was consistently higher in TRI compared to GLU muscle (main effect of muscle: P < 0.05 for all). Transcript L evel of M tDNA E ncoded G enes W as N ot A ffected by A ge As mtDNA content and the expression of mitochondrial biogenesis regulators PGC and TFAM were decreased with age in TRI muscle, we next sought to measure the expression of mtDNA encoded genes ND1 COX1 and COX2 Surprisingly, no age associated changes in any of these mtDNA encoded genes were observed in this study ( P > 0.05 for all; Fig 3A C). When muscles were compared, significant diff erences were detected between GLU and TRI muscle. Specially, the mRNA expression of ND1 COX1 and COX2 was higher in TRI muscle than that of GLU muscle (main muscle effect: P < 0.01 for all), which is consistent with activity and content of CS. Autophagic C apacity W as I mpaired with A ge Along with mitochondrial biogenesis pathways, selective degradation of damaged, dysfunctional mitochondria by (macro)autophagy is considered another

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92 important mechanism to regulate mitochondrial content and homeostasis. To determine whether autophagic flux was impaired with age, we examined the protein expression of nucleoporin 62 (p62, also known as sequestosome 1, SQSTM1; Figure 3 4) in muscle to the autophagosome membrane for subsequent engulfment by the organelle ( Koltai et al. 2012 ), and is finally degra ded in the autophagolysosome (Pankiv et al. 2007 ). We found that protein expression of p 62 was increased in both aged GLU ( P = 0.012) and aged TRI ( P = 0.026) muscle, su ggesting an impairment of autophagic clearance in the aged muscles. No difference was observed between muscles within either young or old horses (main effect of muscle: P = 0.869). Autophagosome F ormation W as I mpacted by A ge Next, we investigated the pos sible causes of the impaired autophagic clearance by analyzing the expression of autophagy related proteins 5 (Atg5) and 7 (Atg7), both essential for autophagosome formation (Jin & Van Remmen, 2009 ). Protein expressio n of Atg5 was reduced in aged TRI ( P = 0.036), and unchanged in aged GLU muscle ( P = 0.266; Figure 3 5D) compared to the respective young counterparts. No age related changes were observed for Atg7 in either muscle (main age effect: P = 0.734; Figure 3 5E) When muscles were compared, TRI exhib ited the same protein expression of Atg5, and higher Atg7 p rotein expression compared to GLU muscle (main muscle effect: P = 0.383 for Atg5, and P < 0.001 for Atg7). LC3 is a well characterized and commonly used autophagosome marker in mammalian cells (Jin & Klionsky, 2013 ). Expression of LC3 mRNA (Figure 3 5C) and protein level of the cy tosolic form of LC3 (LC3 I, Figure 3 5A) were unch anged in aged compared to young TRI muscle ( P > 0.05 for both), while the lip idated form of LC3 (LC3 II, Figure 3 5B), whi ch is integrated in the autophagosomal

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93 membrane and essential for autophagosome formation, was lower ( P = 0.047), further supporting the notion of impaired autophagosome format ion in aged TRI muscle. In the GLU muscle, on the other hand, no aging effects on protein levels of LC3 I or LC3 II were observed despite an age associated increase in t he LC3 transcript level. When GLU and TRI muscles were compared, TRI muscle expressed higher level of LC3 mRNA (main musc le effect: P < 0.001) and LC3 I (main muscle effect: P < 0.05) compared to GLU muscle. Taken together, these findings suggest that the decreased autophagic clearance indicated by elevated p62 levels was at least partially due to impaired autophagosome form ation. Transcript L evel of L ysosomal D egradation M arker LAMP2 W as N ot I mpacted by A ge To further examine whether impaired autophagic clearance could have been caused by defective lysosomal degradation, we determined the mRNA expression level of lysosomal associated membrane protein 2 ( LAMP2 ) using real time qPCR. The protein LAMP2 is needed for efficient fusion of autophagosome and lysosome ( Gonzlez Polo et al. 2005 ), and thereby crucial for completion of the lysosomal autophagic degradation process. Tr anscript levels of LAMP2 (Figure 3 6) did not differ between age groups in either GLU ( P = 0.277) or TRI ( P = 0.592) muscle, implying that decreased autophagic clearance was not due to decreased lysosomal degradation. When muscles were compared, LAMP2 transcript level in TRI m uscle was higher compared to GLU muscle (main muscle effect: P = 0.002). Discussion Mitochondrial damage has been widely discussed for decades as a major contributor to the aging process ( r eviewed by Sun et al. 2016 ). The mitoch ondrial

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94 theory of aging proposes that accumulation of damage to mitochondria and mitochondrial DNA leads to progressive aging in humans and animals (Jang & Van Remmen, 2009 ). Mitochondrial biogenesis and autophagic removal of damaged mitochondria are two p redominate cellular processes whose interplay preserves mitochondrial homeostasis by maintaining mitoc hondrial quality and content (Mishra & Chan, 2016 ). Their functional decline and imbalance could be an underlying cause for cellular aging. Here, we inves tigated how aging affects these two opposing processes, and in conjunction regulate the mitochondrial content in two types of equine skeletal muscle We have previously described the TRI muscle and the GLU muscle in American Quarter Horses as more oxidativ e and more glycolytic, respectively, based on fiber type distribution, citrate synthase activity, and capacity for mitochondrial oxidative phosphorylation. We report here that these muscles with different metabolic properties differed in their response to aging with regard to mitochondrial content. More specifically, a decline in mitochondrial content was only observed in the presuma bly more oxidative TRI muscle, but not in the more glycolytic G LU muscle. In line with decreased mitochondrial content, the decreased gene expression of transcription factors regulating the expression of mitochondrial electron transport system proteins suggests impaired mit ochondrial biogenesis in aged TRI muscle. The combination of age related accumulation of the autophagy car go protein p62 in both muscles and decrease of a utophagosome bound LC3 in the TRI muscle suggest compromised autophagy with age that was more p ronounced in the TRI muscle That this decline in autophagic function was more pronounced is supported by a conco mitant decrease in abundance of the essential autophag y related protein Atg5 in the TRI muscle

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95 Collectively, our results provide the first line of evidence that aging affects mitochondrial biogenesis and autophagy pathways in equine skeletal muscle aging, and that this is likely one of the underlying mechanisms contributing to the observed decrease in mitochondrial content. The decline in mitochondrial content with age has been reported in both humans and animal models ( Barazzoni et al. 2000; Short et al 2005). We (Li et al. 2016 ; Chapter 2 of this dissertation ) and others (Kim et al. 2005 ) have previously shown an age associated decline in mitochondrial content in horse skeletal muscle, indicated by CS activity. In the present study, CS protein conte nt and mtDNA copy number confirmed the changes in mitochondrial content in the TRI muscle we described earlier. Our finding is in agreement with the data reported by others, who showed an age related decline in mtDNA copy number in skeletal muscle ( Welle e t al. 2003 a ; Short et al. 2005 ) from humans, as well as lab animals ( Hartmann et al. 2011; Sczelecki et al. 2014 ). However, other work reported an increased amount of mtDNA in aged skeletal muscle of human ( Pesce et al. 2001 ) and mice ( Masuyama et al. 2005 ). Larsen et al. (Larsen et al. 2012 ) proposed that in contrast to enzyme activity of CS and Complex I of the mitochondrial electron transport chain, mtDNA copy number is a less reliable determinant of mitochondrial content. These authors based thei r conclusion on measurements in skeletal muscle samples from young, healthy humans, but it is conceivable that this poor association of mtDNA copy number with mitochondrial content holds true for aged muscle as well. Furthermore, the strength of this assoc iation might depend on the muscle type investigated, but comparative data are not available.

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96 Mitochondrial DNA copy number can be indicative of mitochondrial biogenesis ( Clay et al ., 2009 ). Mitochondrial biogenesis is a complex process which involves mitochondrial and nuclear DNA encoded genes, recruitment of newly synthesized proteins and lipids, and import and assembly of mitoch ondrial and nuclear products (Zhu et al. 2013 ). We measured mtDNA copy number as surrogate for mtDNA replication in order to evaluate mitochondrial biogenesis in skeletal muscle from aged compared to young horses. Reduced mtDNA copy number obser ved in TRI muscle from aged horses suggeste d an impairment of mitochondrial biogenesis. The transcription factor PGC biogenesis. It associates with the transcription factor NRF 1 to subsequently induce expression of a number of metabolic proteins, including TFA M ( Austin & St Pierre, 2012 ). TFAM translocates to the mitochondria and binds to mtDNA at both the heavy and light strand promoters to not only initiate the transcri ption of mtDNA encoded genes (Fisher et al. 1987 ), but to i nduce m tDNA replication (Bonawitz et al. 2006 ). In accord ance with our findings in the TRI muscle from aged horses, r ecent studies have reported that protein or gene expression of PGC expression was decreased in aged murine ( Koltai et al. 2012; Sczelecki et al. 2014 ) and human skeletal muscle ( Safdar et al. 2010; Ringholm et al. 2013 ). The age related decrease in PGC transcript level in the TRI muscle from aged horses was not concurrent with a lower NRF1 level. However, the mRNA level of TFAM the downs tream target of the PGC NR F1 complex, was lower in aged TRI muscle compared to young horses, which is consistent with the decline in mitochondrial content in this muscle. In contrast to the TRI muscle,

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97 and consistent with mitochondrial content, the more glycolytic GLU muscle displayed no age related alterations of any of the markers of mitochondrial biogenesis assessed in this study. The transcriptional targets of TFAM assessed in this study, which are part of the mitochondrial electron transport system, did not seem to be affected by the apparent age associated dysregulation of the PGC TFAM axis of mitochondrial bio genesis in the more oxidative TRI muscle. The three mitochondrial genes, ND1 COXI and COXII, encode subunits of the mitochondrial electron transport chain: NADH dehydrogenase 1, cytochrome c oxidase subunit 1, and cytochrome c oxidase subunit 2, respectively. The observation that transcription of those genes was not affected by age in either muscl e is consistent with findings in aged rat skeletal muscle ( Barazzoni et al. 2000 ), and suggests that there is a compensatory response for decreased template availability. An increased transcription efficiency to maintain transcript level would help to exp lain the unaltered mitochondrial respiratory capacity observed in a previous study (Li et al. 2016 ; Chapter 2 of this dissertation ). Moreover, we propose that overall ETS activity is more likely to be impaired in individuals older than 80 years ( Lezza et al. 1994; Chabi et al. 2005 ) suffering from severe muscle loss and diseases. Autophagy, also known as cellular self digestion, is a crucial quality control process by which cells sequester and degrade dysfunctional, damaged or aged constituents in order to maintain cellular homeostasis. Mitochondrial autophagy, known as mitophagy, is a selective type of autophagy that specifically eliminates damaged or aged mitochondria ( Ding & Yin, 2012 ). During mitophagy, entire mitochondria are sequestered by an expan ding cup shaped double membrane st ructure, known as

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98 phagophore (Xie & Klionsky, 2007 ). The expansion of the phagophore is regulated by autophagy related, or ATG, proteins, and the phagophore closes and forms a double membrane limited vesicle, the autophag osome. Furthermore, the regulation of autophagosome size by LC3 is critical to allow sequestration of large mitochondria and protein aggregates. Upon completion, the autophagosome fuses with a lysosome, forming the auto(phago)lysosome in which the sequeste red components are ultimately degraded. The impairment of mitophagy was reported to occur in a variety of tissues during normal aging ( Cuervo & Dice, 1998; Donati et al. 2001; Cuervo et al. 2005; Joseph et al. 2013 ). Consequently, an age related accumulation of damaged mi tochondria has been proposed (Terman, 1995 ) and in fact been observed in various cell types of diverse organisms ( Preston et al. 2008; Artal Sanz & Tavernarakis, 2009; Kaeherlein, 2010 ). Consistent with these previous reports, we found that aging was associated with a decreas ed level of autophagy in both GLU and TRI muscles, suggested by p62 protein accumulation. The cargo protein p62 is widely used as a maker of autophagy and involved in mitochondrial aggregation and clearance ( N arendra et al. 2010 ), as it is recruited to mitochondria and targeted for au tophagy mediated degradation (Pankiv et al. 2007 ). Under normal conditions, p62 is rapidly degraded during autophagy, and an impairment of autophagic degradation leads to accumul ation of p62 ( Pankiv et al. 2007 ). Therefore, we propose that the accumulation of p62 in skeletal muscle obse rved by our group and others (Joseph et al. 2013 ) indicates a lower autophagic activity in aged muscle cells. Given the aberrant accumulation of p62 protein in aged skeletal muscles, we further confirmed defects in the autophagic process by analyzing the expression of autophagy related proteins (Atg5, Atg7, LC3

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99 and LAMP2). The protein expression of Atg5, involved in autophagosome formation, signifi cantly declined with age in T RI muscle, but was not significantly affected in GLU muscle. The expression of Atg7, the E1 like enzyme required for autophagosome formation, did not change with age in either muscle, which is in agreement with a previous study in skeletal muscle from old rats (Wohlgemuth et al. 2010 ). Both, Atg5 and Atg7 are essential in the processing of LC3, a homologue of the yeast autophagy protein Atg8. Upon induction of autophagy, LC3 lipidated generat ing a membrane bound form, known as LC3 II. This lipidation process depends on an ubiquitin like conjugation process that involves an Atg12 Atg5 complex, Atg3 and Atg7, which function in concert to facilitate the covalent link of LC3 to autophagosomal prec ursor membranes. Immunoblot analysis showed an age associated decre ase in LC3 II level in equine TRI muscle, which is consistent with some ( McMullen et al. 2009; Carnio et al. 2014; Zhou et al. 2017 ), but not other ( Wohlgemuth et al. 2010; Sebastin et al. 2016 ) studies conducted in other species. It is noteworthy to mention that the lowe r level of LC3 II in the aged TRI muscle measured in our study has likely not resulted from reduced content of LC3 precursor, since neither the level of LC3 mRNA nor L C3 I protein were affected by age. Like LC3 II, expression of Atg5 was also reduced in aged TRI muscle, implying impaired regulation of LC3 lipidation and conjugation to the autophagosomal membrane. No age induced effects were observed in any autophagic ma rkers measured in GLU muscle, suggesting that the formation of autophagosomes, including LC3 lipidation, was not impacted. Overall, these data indicate that upstream autophagy regulators were downregulated in aging equine muscle, and that autophagosome for mation was impaired in aged T RI m uscle,

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100 while autophagy in the GLU muscle appears to be less affected by age. To investigate whether defects of autophagy in equine skeletal muscle extends to processes downstream of autophagosome formation, we assessed expr ession of LAMP2. Lysosomes are the terminal compartment for autophagic degradation, and LAMP2 plays a critical role in the fusion of autophagosomes with lysosomes to form the hybrid structure called autophagolysosome. Reduced fusion has been shown in cells depleted of LAMP2, in which autophagic vacuoles ac cumulated in several tissues (Tanaka et al. 2000 ). We found that gene expression of LAMP2 was not altered with aging in either muscle types, implying that the fusion of autophagosomes with lysosomes may n ot have been impaired at that age. Our findings differ from a previous report in rat skeletal muscle where aging was associated with lower levels of LAMP2 mRNA (Wohlgemuth et al. 2010 ). The divergent response in our study could be attributed to factors in cluding the age of animals, as well as different muscle types being studied. We conclude f ro m our data that the compromised autophagic activity did not appear to be a result of defects in lysosomal involvement. Another noteworthy observation made in our study is the muscle specific response to aging. More specifically, TRI muscle appears more susceptible to th e age related impact than the GLU muscle, indicated by the decrease in mitochondrial content and the more pronounced effect on autophagy related mar kers. We observed here and previously ( Li et al. 2016 ; Chapter 2 of this dissertation ) that the more oxidative TRI muscle contains significantly more mitochondria tha n that of the more glycolytic GLU muscle, independent of age, indicated by CS activity, CS protein content and mtDNA copy number. Similarly, He and coworkers (He et al. 2002 ) reported that the absolute

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101 mtDNA content in single type I oxidative fibers is double that in single type II glyco lytic fibers from human skeletal muscle. In line with the higher mitoc hondrial content we observed, TRI muscle also displayed a higher expression level of mitochondrial genes ( ND1 COXI and COX II ), mitochondrial biogenesis markers ( PGC NRF1 and TFAM ) and higher proteins levels of autophagy regulators (Atg7, LC3, LC3 I, and LC3 II) when compared to GLU muscle. Inte restingly, the more oxidative TRI is also more likely to suffer an age related decline in mitochondrial content co mpared to the more glycoly tic GLU Similarly, in rodents CS activity declined with age in the oxidative soleus but not in the more glycolytic extensor digitorum longus ( Picard et al. 2011 a ), and mtDNA content decreased with age only in soleus and tibialis anterior but not in gast rocnemius or extensor digitorum longus muscles (Pesce et al. 2005 ). In addition, differential effects of aging on CS activity were observed in human skeletal muscle, with CS activity declining in the gastrocnemius but not in the vastus lateralis These results suggest that the age associated decline in mitochondrial content in skeletal muscle is not uniform. However, it appears to follow a common pattern. The decline in mitochondrial content is larger and begins at an earlier age in muscles with high oxi dative capacity ( soleus TRI ) compared to less oxidative mixed muscles ( gastrocnemius; GLU ); and muscles with high glycolytic capacity ( extensor digitorum longus and vastus lateralis ) appear to be less impacted than these other muscle types. The response o f the tibialis anterior described by Pesce et al. (Pesce et al. 2005 ) seems to represent an exception to this trend, highlighting the lack of uniformity of the age effects in skeletal muscle. It might reflect a differential use of these muscles with aging affecting the intracellular response, but this context has not yet been illuminated.

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102 The differences in susceptibility to aging between the two muscles observed in our study could be attributed to the differential regulation of mitochondrial biogenesis a nd autophagy. The imbalance between mitochondrial degradation and biogenesis may underlie the age associated decline in mitochondrial content. Maintenance of a stable pool of healthy mitochondria requires both autophagy and mitochondrial biogenesis. Howeve r, impairment in both biogenesis and autoph agy was observed with age in TRI muscle, and decreased mitochondrial content suggests that the effects of declined mitochondrial biogenesis exceeded that of declined autophagy. While we also observed an impairment of autophagic activity in the GLU muscle from aged horses, neither mitochondrial content nor biogenesis were affected in this muscle. It is possible that a more severe imbalance of mitochondrial biogenesis and degradation commences a t a later age compared to the TRI muscle. In summary, we report here, for the first time, the age associated alterations in mitochondrial biogenesis and autophagy in equine skeletal muscle. We observed a decreased mitochondrial content wit h age in TRI muscle but not in GLU muscle, implying that the age related decrease in mitochondrial content is muscle type specific. With age, mitochondrial biogenesis and autophagy were impaired, which might be one underlying cause of the age associated decline in mitochondrial content. The se findings suggest that a targeted approach to balance mitochondrial biogenesis and degradation may prolong health and performance of aged American Quarter Horses.

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103 Table 3 1. Primers used for gene amplification in q uantitative reverse transcription polymerase chain reaction Gene GenBank Accession no. Product size (bp) PGC XM_014738763.1 GAACAGCAGCAGAGACAAATG GGGTCAGAGGAAGAGACAAAG 104 NRF1 XM_014739148.1 GTGGTCCAGACCTTTAGTAACC CCATCAGCCACAGCAGAATA 146 TFAM XM_001503382.3 CTCAGAACCCAGATGCGAAA CTGCCCTGTAAGCATCTTCATA 108 LC3 XM_005608485.2 CTCAGGAGACATTTGGGATGAA CGGATCGATCTCAGTTGGTAAC 119 LAMP2 XM_014729146.1 TGAACGTCACTCACGATAAGG AGCCTAAGTAGAGCAGTTTGAG 100 COX2 NC_001640.1 TCATCCGAAGACGTCCTACA GCCACGAGAGTTGTCTGATTTA 95 COX1 NC_001640.1 CAGACCGTAACCTGAACACTAC GGGTGTCCGAAGAATCAGAATAG 91 ND1 NC_001640.1 GGATGGGCCTCAAACTCAA GGAGGACTGAGAGTAGGATGAT 106 ACTB NM_001081838.1 CTCCATTCTGGCCTCATTGT AGAAGCATTTGCGGTGGA 98 GAPDH NM_001163856.1 GTCATCATCTCTGCTCCTTCTG GGAGGCATTGCTGACAATCT 99

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104 Figure 3 1 Protein expression of citrate synthase in skeletal muscle from American Quarter Horses. Densitometric quantification of citrate synthase (CS) protein expression in G LU and TRI muscle from young (n = 24 for GLU, n = 12 for TRI) and aged (n = 12 for GLU, n = 12 for TRI) horses. Values are mean s SE. Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.05. GLU vs. TRI: §§ P < 0.01. Representative Western blot images are shown above the graph.

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105 Figure 3 2. Mitochondrial DNA copy number and transcript levels of factors associated with mitochondrial biogenesis in skeletal muscle from American Quarter Horses. Mitochondrial DNA copy number from GLU and TRI muscle of young (n = 7 for GLU, n = 8 for TRI) and aged (n = 6 for GLU, n = 6 for TRI) horses was determined using quantitative real time PCR of ND1 (A) and COX1 (B), actin gene ( ACTB ). C) Correlation between mtDNA copy number ( COX1 / ACTB ) and citrate s ynthase pr otein expression in TRI and G LU muscle (insert : GLU ). Each point represents an individual muscle sample from young (open symbols) and aged (closed symbols) subjects. D F) Real time PCR analysis of PGC NRF1 and TFAM mRNA levels in GLU and TRI m uscle fro m young (n = 7 8 for GLU, n = 8 f or TRI) and aged (n = 7 8 for GLU, n = 8 for TRI) horses. Results are represented as the fold change compared to young GLU muscle. Values are mean s SE. Open bars represent young horses; solid bars, aged horses Young vs. aged: P < 0.05 GLU vs. TRI : § P < 0.05, §§§ P < 0.001.

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106 Figure 3 2. Continued

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107 Figure 3 3 Transcript level of mtDNA encoded genes in skeletal muscle from American Quarter Horses. Expression of ND1 (A), COX1 (B), and COX2 (C) mRNA in GLU and TRI muscle from young ( n= 7 8 for GLU, n = 8 for TRI ) and aged (n = 7 8 for GLU n = 7 8 for TRI ) horses. Results are represented as the fold change compared to young GLU muscle. Values are mean s SE. Open bars represent young horses; solid bars, aged horses. GLU vs. TRI : §§ P < 0.01, §§§ P < 0.001.

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108 Figure 3 4. Protein expression of p62 in skeletal muscle from American Quarter Horses. Protein level of p62 in GLU and TRI muscle from young (n = 24 for GLU, n = 12 for TRI ) and age d (n = 12 for GLU, n = 12 for TRI ) horses. Representative Western blot images are shown above the summary graph. Values are mean s SE. Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.05.

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109 Figure 3 5 Gene and protein expression of autophagy regulatory proteins in skeletal muscle from American Quarter Horses. Protein expression of LC3 I (A) and LC3 II (B), and mRNA expression of LC3 (C); and protein expression of Atg5 (D) and Atg7 (E) in GLU and TRI mu scle from young ( mRNA expression: n = 8 for GLU, n = 8 for TRI ; protein expression: n = 22 24 for GLU, n = 11 12 for TRI ) and aged ( mRNA expression: n = 8 for GLU, n = 8 for TRI ; protein expression: n = 12 for GLU, n = 11 12 for TRI) horses. F) Representative Western blot images of Atg5, Atg7 and LC3 I and LC3 II in GLU and TRI muscle from young and aged horses. Values are mean s SE. Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.05, *** P < 0.001. GLU vs. TRI : § P < 0.05, §§§ P < 0.001.

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110 Figure 3 5. Continued

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111 Figure 3 6 Transcript level of the LAMP2 gene in skeletal muscle from American Quarter Horses Expression of LAMP2 mRNA in GLU and TRI muscle from young (n = 8 for GLU; n = 8 for TRI ) and aged ( n = 8 for GLU; n = 8 for TRI ) horses is represented as the fold change compared to young GLU muscle. Values are mean s SE. Open bars represent young horses; solid bars, aged h orses. GLU vs. TRI : §§ P < 0.01.

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112 CHAPTER 4 AGE RELATED CHANGES IN MYOGENIC CAPACITY OF SATELLITE CELLS OBTAINED FROM AMERICAN QUARTER HORSES Background Healthy skeletal muscle has a remarkable capacity to repair injury via muscle regeneration. In adults, this ability is owed to muscle stem cells, commonly referred to as satellite cells. When those cells were first identified in 1961 (Mauro 1961), they were ma membrane of myofibers. Given the fact that they represent the only source of new myonuclei in postnatal skeletal muscle, satellite cells might be centrally involved in maintenance of healthy muscle mass (Biressi & Rando, 2010). It is hypothesized that a decline in satellite cell function and/or number is a main factor causing the insufficiency of muscle regeneration in elderly humans, which subsequently leads to a loss of myofibers and a decline in muscle cross sectional area, recognized as sarcopenia (G opinath & Rando, 2008; Garca Prat et al. 2013). Moreover, a number of studies have shown type II muscle fiber atrophy with aging is accompanied by a fibertype specific decline in satellite cell content ( Verdijk et al. 2007; Verney et al. 2008; McKay et al. 2012). In general, satellite cell content and function are suggested to decrease with age (Conboy et al. 2003; McKay et al. 2013; Fry et al. 2015 ), rendering muscle regeneration insufficient in aged muscle ( Brooks & Faulkner, 1990; Ikemoto Uezumi et al. 2015 ). However, these findings were based on studies, in which satellite cell regenerative capacity was tested in vivo in response to physiological and pathological stimuli, including exercise and injury. The effects of aging on satellite cell s the mselves were not examined directly.

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113 Although the exact underlying mechanisms contributing to satellite cell dysfunction are not yet fully understood, increasing evidence gathered over the past decade showed that diminished satellite cell function is due t o both intrinsic changes in satellite cells themselves and extrinsic alterations in the satellite cell microenvironment, also called the satellite cell niche. Support for the latter mainly comes f ro m transplantation experiments, in which declined regenerat ive capacity of old satellite cells ( Conboy et al. 2005; Villeda et al. 2011; Lavasani et al. 2012 ), while young satellite cells lost their regenerative function when Carlson & Faulkner, 1989 ). Intrinsic changes of satellite cell s with age may also regulate satellite cell function. In particular, over the past a few years, the role of mitochondria for satellite cell performance has been increasingly appreciated. As expected given their essential role for energy production, among other vital functions ( Finkel & Holbrook, 2000; Ryan & Hoogenraad, 2007 ), mitochondrial function appears to be a major player in regulating satellite cell myogen ic properties ( Marzetti et al. 2013; Siegel et al. 2013) Mitochondrial function has been suggested essential for satellite cell maintenance and activation ( Cerletti et al. 2012; Stein & Imai et al. 2014; Ryall et al. 2015 ). In addition, an increase in mitochondrial respiratory function is indispensable during differentiation from myoblasts into myotubes in order to establish the oxidative phenotype that is necessitated by the high energetic demand of muscle contraction. Upon differentiation, myoblasts shift from a previous glycolytic state to predominant reliance mainly on mitochondrial ox idative phosphorylation This shift is accompanied by a marked increase in mitochondrial biogenesis and mitochondrial oxidative respiration ( Remels et

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114 al. 2010 ). However, mitochondrial function is impaired in old satellite cells, indicated by downregulation of the TCA cycle and OXPHOS reactions (Kuilman et al. 2010; Lpez Otn et al. 2013; Zhang et al. 2016). Mitochondrial quality control mechanisms, including mitochondria specific proteases and chaperones, fission/fusion processes, mitochondrial biogenesis and autophagy, are responsible for maintenance of mitochondrial homeostasis, and therefore, might also play an important role in regula ting mitochondrial function in satellite cells. Indirect support for the role of mitochondria in satellite cell function comes from studies in which satellite cell function was altered in conditions that improved or harmed mitochondrial homeostasis. For example, short term caloric restriction, which is known to stimulate mitochondrial biogenesis, stimulated satellite cell proliferation in young and old mice (Cerletti et al. 2012), and alterations in mitochondrial biogenesis led to failure of satellit e cell induced muscle regeneration during conditions of muscle wasting (Toledo et al. 2011). Further, impaired autophagy led to increased ROS generation, which in turn drove satellite cell senescence in mice (Garca Prat et al. 2016), while autophagy rea ctivation ameliorated muscle regeneration in mdx mice (Fiacco et al. 2016). Horses are among the most athletic animals, and they may suffer subtle muscle injuries on a regular basis during their normal daily activities ( Taylor et al. 1981). Continuous m therefore essential for muscle mass maintenance throughout life. However, not much is known about the effects of aging on satellite cell function in equine muscle. Furthermore to the best of our knowledge, no studies have so far investigated age related alterations in mitochondrial quality control, pivotal in maintaining mitochondrial

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115 function, in horse satellite cells; and it is worth noting that those control mechanisms may be negatively affected during the aging process. Therefore, the overall goal of this project w as to examine differences in the intrinsic myogenic capacity and mitochondrial quality control mechanisms of satellite cells isolated from young and aged horses, and to compare different muscle types. Materials and Methods Animals and M uscle S ample C ollection Clinically healthy young (n = 4, 1 gelding and 3 mares, aged 2 4 years) and aged (n = 4, 1 gelding and 3 mares, aged 20 27 years) American Quarter Horses were used in this study. All horses were owned by the University of Florida and held on pasture at received forced exercis e for 6 mo prior to the study. All procedures performed in this study were approved by the University of Florida Institute of Food and Agricultural Sciences Animal Research Committee. Skeletal muscle microbiopsies were collected in a sterile manner from th e GLU and TRI following the procedure described in Li et al. ( Li et al. 2016 ; Chapter 2 of this dissertation ). Briefly, following administration of local anesthesia, muscle samples were collected at a sampling depth of 5 cm, using a 14 gauge SuperCore TM Biopsy needle (Angiotech, Gainesville, FL, USA). Muscle samples were placed in ice cold Dulbecco's phosphate buffered saline (DPBS, Hyclone TM South Logan, UT, USA) and immediately transported to the lab for satellite cell isolation. As a comparison of cel ls from young and aged horses was required, muscle was removed from young and aged horse sequentially and processed in parallel throughout all experiments.

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116 Satellite C ell I solation Under sterile conditions, muscle samples were washed with PBS to remove an y surface blood. After re suspend ing in fresh PBS, visible fat and connective tissue were cleaned off, and the remaining tissue minced into a coarse slurry and subsequently digested using pronase (Sigma Aldrich, St. Louis, MO) for 1 h at 37 C The digested tissues were then centrifuged at 1600 g for 10 min, the pellets were re suspended in PBS and filtered through a 100 another centrifugation at 1600 g for 10 min, cells were re susp ended in growth 10 (Corning, NY) supplemented with 20% horse serum (Gibco Thermo Fisher Scientific, Waltham, MA, USA), 1% L glutamine (Corning), and 1% Antibiotic Antimycotic (Gibco Thermo Fisher Scientific), plated on 100 mm cultu re dishes coated with gelatin (Sigma Aldrich), and cultured in an incubator with 5% CO 2 at 37 C. Media were changed every 48 h. Cells were passaged on day 7 after isolation and subsequently every 2 days (at 60 70% confluence). Cell were passaged three tim es in total and then cryopr eserved until further use. A 30 min pre plate was performed at each passage step to enrich satellite cells population. Proliferation and D ifferentiation A ssays After thawing cryopreserved cells (as passage #4), they were cultured in GM, with medium exchange every 24 h. Cell were trypsinized when they reached 60 70% confluency, washed in PBS once and this 5 th cell passage divided into experimental batches as outlined below. At this 5 th passage, cultures contained >90% myog enic cells, based on the express ion of the myogenic marker, Pax 7, analyzed by immunofluorescent staining using anti Pax7 (1:2, Developmental Studies Hybridoma

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117 Bank (DSHB), Iowa City, IA, USA) and Alexa fluor 488 conjugated secondary rabbit anti mouse IgG ( 1:500) (see also section 2.4). For differentiation analysis, cells of the 5 th passage were allowed to proliferate in GM until they reached 80 90% (submaximal) confluency. At that time, the GM was exchanged for differentiation media (DM), DMEM with low glu cose (Corning) supplemented with 1% sodium pyruvate (Lonza, Walkersville, MD ), 1% HEPES (Gibco ), 1% L glutamine (Corning), 1% insulin transferrin selenium (Gibco ), and 1% Anti Anti (Gibco ). The differentiation media induces satellite cell differentiatio n due to the low serum content. Cells were cultured in DM for the indicated time (24 h, 48 h, and 96 h). Because cells from aged horses exhibited a lower proliferation rate in practice experiments, they were prepared and seeded 5 h before cells from young horse s in order to reach submaximal confluency at the same time as cells from young horses for simultaneous performance of the differentiation assays. Measurement of M yoblast D ifferentiation and F usion Cells were seeded at a density of 4 10 3 /well in 96 well plates in quadruplicate, and the GM exchanged for DM when t he cells reached 80 90% confluency (D0 DM, day 0 in DM) as described above. On day 3 in DM (D3 DM), following a quick rinse with PBS, cells were fixed and permeabilized in 100% met hanol for 10 min at 20 C, and subsequently blocked with 5% horse serum in PBS for 45 min at room temperature (RT). Cells were then immunostained with primary antibodies for 1h at RT followed by an overnight incubation in primary antibody at 4C. Primary antibodies used were as follows: myogenin (2 wells/sample; 1:2, F5D, DSHB) and myosin heavy chain (MyHC) (2 wells/sample; 1:2.4, MF20, DSHB). After incubation with primary antibodies and a quick rinse with PBS supplemented with 0.5% BSA (Sigma Aldrich) cel ls were

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118 incubated with Alexa fluor 488 conjugated secondary rabbit anti mouse IgG (1:500 for MyoG; 1:1000 for MyHC, Thermo Fisher Scientific) for 1h at RT. All immunofluorescently labeled cells were counterstained with Hoechst 33342 (1g/mL, Thermo Fisher S cientific) to label the nuclei. Cells were examined with an EVOS fluorescence microscope (Thermo Fisher Scientific), and labeled cells manually counted and recorded using ImageJ software. Myogenin (a marker of cell differentiation) staining was expressed a s a percentage of myogenin labeled cells relative to the total number of cells (labeled by nuclear stain) in the field of view The efficiency of fusion (fusion index) was determined by counting the number of nuclei in MyHC positive myofibers as a percenta ge of the total number of nuclei in the field of view. At least five randomly selected fields of view were imaged per sample. The mean of these measurements was taken as the sample value. Isolation of T otal RNA and Real T ime qPCR Total RNA was isolated from differentiating cells (D1 DM and D2 DM) using TRIzol (Thermo Fisher Scientific), with genomic DNA removed using a DNA free TM Kit Immediately following DNA removal, RNA concentration and purity were determined by measuring the absorbance at 260 and 280 nm with a UV spectrophotometer (Synergy HT, BioTek Instruments ). Two microgram s of total RNA was reverse transcribed in a 20 ion system performed with a High Capacity cDNA Reverse Transcription Kit (Applied Subsequently, PCR amplification was carried out using SYBR green fluorescence ( iTaq TM Universal SYBR Green Supermix Bio Rad) on a CFX Connect real time PCR detection system (Bio Each assay plate

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119 contained negative controls and a relative standard curve generated with five serial 5 fold dilutions of cDNA pooled from experimental samples. The standard curve was accepted at a R 2 of 0.99 or greater, otherwise the samples were repeated. Melting curve analyses were performed to verify the amplification of a single PCR product. Transcript levels were ana lyzed using relative standard curve, normalized to GAPDH mRNA, and results were expressed as a fold change in mRNA level compared with expression in GLU from the young horse group. All reactions were performed in duplicate. Specific primer sequences are listed in Table 3 1. MtDNA C opy N umber M easurement Total DNA was extracted from differentiating myoblasts (D1 DM and D2 DM) using Wizard Genomic DNA purification kit (Promega Corporation, Madison, WI) determined using a UV spectrophotometer ( Synergy HT, BioTek Instruments ). Relative quantification of mtDNA copy number was performed by real time qPCR using primers specific for mitochondrial gene ND1 ( NADH dehydrogenase 1, forward: GGA TGG GCC TCA AAC TCA A GGA GGA CTG AGA GTA GGA TGA T ) and normalized against the nuclear gene ACTB ( actin forward: CTC CAT TCT GGC CTC ATT GT ; reverse: AGA AGC ATT TGC GGT GGA ). To confirm the results assessed with ND1 primers, another pair of primers designed within mitochondria gene COX1 ( cytochrome c oxida se 1 ) region was also tested. Primer sequences for COX1 were provided below: CAG ACC GTA ACC TGA ACA CTA C GGG TGT CCG AAG AAT CAG AAT AG DNA was amplified using a CFX Connect real time PCR detection system (Bio Rad) wi th conditions listed below: 95 C for 3 min and 40 cycles of 95 C for 5 seconds and 60 C for 30 seconds. Melting curve

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120 analysis was performed from 65 C to 95 C. The relative quantification was done using the relative standard method (five serial dilut ions of pooled DNA samples) and the R 2 of mitochondrial gene to genomic single copy gene, resulting in a mtDNA/nDNA ratio, a value proportional to the average number of mtDNA copy number in each sample. All s amples were amplified in duplicate on the same plates. Immunoblo t ting Whole cell lysates (D1 DM and D2 DM) were obtained by applying Pierce TM RIPA buffer (Thermo Fisher Scientific) containing 1% Halt TM protease phosphatase inhibitor cocktail (Thermo Fish er Scientific) to adherent cells, followed by scraping the lysed cells off the substrate. Cell lysates were stored at 80 C until further analysis. Proteins were quantified using Thermo Scientific TM PierceTM BCA protein assay kit (Thermo Fisher Scientific ). Equal amounts of the total protein were electrophoretically separated on a Sodium dodecyl sulfate polyacrylamide gel (BioRad) and transferred to a Polyvinylidene fluoride membrane (PVDF; Thermo Fisher Scientific), and immunoblotted under standard condit ions as previously described (Wohlgemuth et al. 2010). Primary antibodies used were as follows: anti p62/SQSTM1 (1:300, Sigma Aldrich), anti CS (1:100, Santa Cruz Biotechnology, Dallas, TX), anti Hsp60 (1:1000, Cell Signaling Technology, Danvers, MA), ant i LC3 ( 1:500, Thermo Fisher Scientific), and anti tubulin (1:20,000, Sigma Aldrich). Secondary antibodies were either horseradish peroxidase or alkaline phosphatase conjugated (Sigma Aldrich). Membranes were then developed with DuoLux chemiluminescent/fl uorescent substrate (Vector Laboratories, Burlingame, CA) and the chemiluminescent signals captured with a digital imager (G:Box Chemi XR5, Syngene). Densitometric analysis of band signals was performed using the

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121 s, Syngene). Protein level was expressed relative to tubulin expression, which served as a loading control. Statistical A nalysis Dat a are pre sented as means SE with the number of samples per group indicated in the figure legends. Statistical analysis was performed using SigmaPlot 13.0 software (Systat Software Inc., San Jose, CA). For data that were not normally distributed as determined by Shapiro Wilk normality test, a log 10 transformation was performed in order to achieve a normal distribut ion Data were analyzed for significance by two way analysis of variance with factors of age ( Young Aged ) and muscle type ( GLU TRI ), followed by posthoc Holm Sidak multiple comparison tests. Data were considered significantly different if the P value 0.05. Trends were reported if 0.05 < P < 0.1. Results Satellite C ells from A ged H orses S how ed R educed M yogenic P otential To measure the effect of aging on satellite c ell proliferative capacity (Figure 4 1), cells were grown in GM for 6 days until the cells reached confluency. Cells were harv ested each day from day 3 (D3) in GM, and the number of cells per well counted. In general, cell number progressively increased from D3 to D6 regardless of age of origin and of muscle type. However, wells with cells obtained from aged TRI tended to have a lower cell number compared to those with cells obtained from young TRI on D3 ( P = 0.086). Similarly, on D4, wells with aged TRI cells tended to have fewer cel ls compared to those with young TRI cells ( P = 0.062), wh ile significant lower cell numbers were observed in wells with a ged GLU cells compared to young GLU cells on D4 ( P = 0.015). After D5, young and aged cells showed similar proliferative capacity, independent of

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122 muscle type. To determine changes in satellite cell differentiation capacity, cells were labeled with myogenin (a key marker of differentiation) and MyHC (a marker for terminal differentiation) on day 3 after induction o f differentiation (D3 DM) The number of myogenin + cells ( Figure 4 2) decreased with age (main age effect: P = 0.007). More specifically, the percentage of myogenin + cells (Figure 4 3) was decreased by 33% in cells from aged TRI ( P = 0.017), and ~30% in cells from aged GLU ( P = 0.079). Consistent with these findings, the percentage of nuclei in MyHC + cells was lower in ce lls from aged compared to young TRI. However, the fusion capacity (fusion index, number of nuclei in MyHC + cells/ total number of nuclei in the field of view) was similar between ce lls derived from young a nd aged GLU. These data suggest that satellite cells obtained from aged horses have impairments in proliferation and differentiation, and TRI derived satellite ce lls are more susceptible to age related changes. Mitochondrial DNA C opy W as E levated with A g e Differentiation of satellite cells (myoblasts) into myotubes requires a metabolic switch from glycolysis to OXPHOS. In order to meet the energy demands of myotubes, the myoblasts must generate sufficient mitochondria before they fuse into myotubes (day 2 postdifferentiation). To investigate whether mitochondrial density differs between age groups on D1 DM and D2 DM, the protein expression of CS a key mitochondrial matrix enzyme and commonly used as a marker for mitochondrial content (Larsen et al. 2012), was analyzed (Figure 4 4 ). No age related changes in CS content were observed in cells from either muscle type at either time point ( P > 0.1 for all). Surprisingly, mtDNA copy number analyzed using ND1 primers was increa sed in aged cells at D2 DM (F igure 4 5 ; P = 0.056 for GLU, P = 0.033 for TRI), but did not d iffer at D1 DM (Figure 4 5 ; P > 0.1 for all). This was also the case when primers of another

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123 mitochondrial gene, COX1, was used, which confirmed that mtDNA copy number was elevated with age in both GLU and TRI derived cells at D2 DM ( P = 0.028 for GLU; P = 0.021 for TRI). Taken together, our data indicate that the mtDNA copy number is elevated in both GLU and TRI derived satellite cells form aged horses. Mitochondrial G enes W ere D ownregulated in D ifferentiated C ells from A ged TRI M uscle Given the elevated mtDNA copy number in cells from aged muscles, changes in mRNA level of the mtDNA encoded genes ND1 COX1 and COX2 was determined using real time qPCR. The increase in mtDNA copy number did not cause a concomitant increase in transcript level o f these mtDNA encoded genes. In fact, gene expression of ND1 was reduced by 63% in ce lls from aged TRI at D1 DM (Figure 4 6 P = 0.007), and this reduction was even pronounced, ~80%, at D2 DM (Fig ure 4 6 P = 0.043). No apparent age associated changes in ND1 gene expression were detected in cells derived from aged GLU at either time point. Similarly, COX1 expression of cells from aged TRI decreased by 43% at D1 DM ( P = 0.031), and by 64% at D2 DM ( P = 0.01 7) compared to cells from young TRI. No age associated changes in COX1 gene expression were observed in cells derived from aged GLU at any point during differentiation. Gene expression of COX2 did not differ between age groups at either D1 DM or D2 DM cells from in either muscle. Collectively, our gene expression data demons trate that transcript levels of mitochondrial genes are downregulated in differentiated cells derived from aged TRI, which might indicate an impairment of mitochondrial OXPHOS capacity with age.

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124 Satellite C ells D erived from A ged M uscle E xhibited I mpaired M itochondrial Q uality C ontrol M echanism To investigate possible mechanisms underling the impaired satellite cell regenerative capacity observed in this study, markers relevant to mitochondrial quality control systems were assessed. Protein expression of a mitochondrial specific chaperone, Hsp60, as determined in diff erentiated satellite cells (Figure 4 7 ). No significant difference in Hsp60 content was observed between cells derived from young and aged horses in either muscle, suggesting that cells maintain their ability to regulate protein foldi ng and refolding through Hsp60 with age. Gene expression analysis of mitochondrial biogenesis markers identified a decreased transcription of PGC ( P = 0.014; Figure 4 8 ), but unaltered NRF1 and TFAM (Figure 4 8 ) in satellite cells obtained from aged T RI on D1 DM. On D2 DM, the decrease in PGC expression was less pronounced ( P = 0.059; Figure 4 8 ), while gene expression of NRF1 ( P = 0.05; Figure 4 8 ) and TFAM ( P = 0.008; Figure 4 8 ) was lower in satellite cells deri ved from aged compared to young TRI at that time point. For cells derived from GLU, expression of all markers relevant to mitochondrial biogenesis investigated here did not change with age on D1 DM (Figure 4 8 ). However, there was a dramatic decline in PGC expression on D2 DM in cells from aged GLU ( P = 0.024; Figuire 4 8 ), which was not accompanied by any change in expression of NRF1 and TFAM (Figure 4 8 ). Collectively, these findings suggest that mitochondrial biogenesis was impaired with age in differentiated satellite cells. To investigate autophagic activity during satellite cell differentiation, the autophagy regulators p62, LC3 and LAMP2 were assessed using either western blot or real time qPCR, or both. At D1 DM, accumulation of p62 ( P = 0.038) suggests an

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125 impairment of autop hagic degradation (Figure 4 9 ) in cells obtained from aged TRI. Consistent with this finding, an elevated level of the autophagosomal membrane bound, phosphatidylethanolamine conjugated form of LC3 (LC3 II; P = 0.034; Figure 4 10 ) was detected in this grou p. To investigate whether the downstream process of fusion of the autophagosome with the lysosome was affected by age, expression of LAMP2 was examined (Figure 4 11) Transcript level of LAMP2 seemed to be numerically lower in cells obtained from aged TRI, but it the variance was high and the difference to cells derived from young TRI did not reach significance. At D2 DM, cells from aged TRI continued to exhibit an accumulation of p62 ( P = 0.048). Moreover, there was a dramatic reduction in LC3 gene express ion (by 76%, P = 0.011) with age in TRI cells. Neither LC3 I or LC3 II protein level differed between cell from young and aged TRI. Cells obtained from aged GLU did not show significant changes in expression of either autophagy regulator, but there was a tendency for a decreased LC3 II protein level ( P = 0.076). Our data indicate poor autophagic degradation activity in differentiated cells obtained from aged TRI. Discussion It has been suggested that aged muscle exhibits poor repair and regeneration capacity following exercise induced damage ( Brooks & Faulkner, 1990). However, knowledge on satellite cell biology in healthy aging objectes is still insufficient. Especially in equine, it is largely unknown whether satellite cells of old horses differ in their intrinsic myogenic capacity from those of young individuals. Among the hallmarks of aging, mitochondrial dysfunction has been in the spotlight for a long time. The maintenance of a functional mitochondrial pool is essential for cellular processes, in particular for muscle regeneration where differentiated cells are strictly dependent on

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126 oxidative, mitochondria supported metabolism (Wagatsuma & Sakuma, 2013). Indeed, given the role of mitochondria in cell differentiation, the question arises whether ch anges in mitochondrial density and/or function, or alteration in mitochondrial quality control pathways, could subsequently affect satellite cell myogenic capacity. This work is the first to compare the intrinsic myogenic potential of satellite cells isol ated from healthy young and aged horses, and to provide evidence that impaired mitochondrial quality control mechanisms could be a potential cause of the failure of proper satellite cell differentiation. Primary culture of satellite cells isolated from mus cle biopsies under standard conditions in vitro showed that cells derived from aged horses tended to have a lower cell proliferation rate and significantly reduced differentiation capacity compared to those derived from young horses. Moreover, these age as sociated impairments in cell myogenic capacity were more prono unced in cells from the TRI muscle than those from the GLU muscle. Noteworthy, reduced myogenic potential of aged cells was accompanied by an elevation of mtDNA copy number without a concomitant increase in mitochondrial density and function. What is more, the protein content of citrate synthase, a marker of mitochondrial density (Larsen et al. 2012), did not change with age, and the transcript levels of mtDNA encoded genes were downregulated in aged cells, indicating impaired mitochondrial function. The age dependent decline in gene expression of mitochondrial biogenesis markers ( PGC NRF1 and TFAM ) and the alterations in some of the autophagy markers (p62 and LC3) in aged TRI cells point to insufficient mitochondrial quality control as one of the underlying causes of mitochondrial dysfunction, which in turn may lead to compromised myogenic capacity. At present, our data do not sufficiently explain why the myogenic

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127 potential of cells obtained from aged GLU were slightly impacted, but the significant reduction in PGC and a tendency for decreased LC3 II protein could point in the same direction. It is possible, that the more glycolytic GLU experiences the negative impact of aging on satellite cell function at a later age compared to the more oxidative TRI, but this speculation warrants further evaluation. Skeletal muscle regeneration depends greatly on the interplay between the satellite cell and its local microenvironment, known as the satel lite cell niche. In most of the injury induced muscle regeneration models, the muscle regenerative outcomes seemed to depend more on the age of the niche, rather than on the aged satellite cell itself (Grounds, 1998). Following injury, a dramatic change in levels of growth factors and cytokines has been observed within the satellite cell niche ( Edwall et al. 1989; Kurek et al. 1996; Warren et al. 2002). Aged horses, for example, exhibited increased expression of inflammatory cytokines IL 6, and IF in circulation (McFarlane & Holbrook, 2008), which have been reported to impede cell proliferation and differentiation of C2C12 myoblasts (Al Shanti & Stewart, 2012) as well as horse satellite cells in vitro (LaVigne et al. 2015). In addition, aged satellite cells failed to respond to repair stimuli, such as induced injury (Conboy et al. 2003; Gopinath & Rando 2008; Kuang et al. 2008 ), due to a decline in satellite cell activation (Carlson et al 2008; Conboy et al 2003). It is important to note that these extrinsic alterations of the niche, such ascytokine levels, and the distinct sensitivities to repair stimulation signals could mask intrinsic changes in the aged satellite cell itself. Furthermore, those experiment ally induced or extreme types of injuries are different from physiological attrition and muscle regeneration stimulated by rout ine physical activities. Our results

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128 point to intrinsic changes with age and thereby can explain, at least in part, why the regen erative capacity is reduced in old compared to adult satellite cells exposed to the same environment ( Sousa Victor et al. 2014 ), and why old satellite cells can be rejuvenated by specific molecular cues (Carlson et al. microenvi ronment ( Carlson & Conboy 2007 ). One finding of the present study was the difference in satellite cell susceptibility to aging between the two muscles. Satellite cells isolated from TRI were more susceptible to age when compared to cells isolated from GLU. More specifically, TRI derived a ged cells failed to differentiate as well as young cells, which was concomitant to dysregulation of mitochondrial biogenesis and autophagic degradation pathways, and probably consequential diminished mitochondrial function. We have previously shown tha t the fibertype composition of the TRI is more oxidative than that of the GLU (Li et al. 2016 ; Chapter 2 of this dissertation ). At the applied sampling depth, TRI contained ~10% MyHC I and 35% MyHC IIA, and the GLU ~3% MyHC I and 21% MyHC IIA, which is si milar to data published by van den Hoven et al. ( van den Hoven et al. 1985), who reported a higher percentage of type I fibers in TRI compare to GLU in Dutch Saddle Ho rses at the same sampling depth. Satellite cell number and function differ between muscl e types. For example, t he percentage of satellite cells in rat soleus muscle (predominan tly type I fibers) is around 2 fold higher than in more glycolytic muscles, such as the tibialis anterior or extensor digitorum longus muscle (Gibson & Schultz, 1982; Okada et al. 1 984). The different respo nse of intrinsic satellite cell function s to aging could be explained by a different level of oxidative stress the cells might be exposed to. During aging, the production of ROS was elevated within

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129 my ofibers (Chabi et al. 2008), which can subsequently cause oxidative damage to surrounding structures, including satellite cells, which are believed to have reduced antioxidant capacity with aging in human (Fulle et al. 2005). The more oxidative TRI is su rmised to exert more oxidative stress and damage, due to its higher OXPHOS activity and presumably a concomitantly higher ROS, and to its greater use in routine activities (posture) compared to the GLU (more voluntary and propulsive activities). Consequent ly, its satellite cells might experience a higher level of oxidative stress and damage, which could underly progressive dysfunction with age. However, this hypothesis needs to be tested further. Stem cells are known to contain few mitochondria and rely greatly on glycolytic metabolism for energy production, while when differentiated they reportedly rely mostly on OXPHOS for their energy demand (Leary et al. 1998; Wagatsuma & Sakuma, 2013). In sup port, a dramatic remodeling of the mitochondrial network has been observed during stem cell differentiation (Sin et al. 2016). Interestingly, an impaired mitochondrial network during differentiation is suggested to negatively affect muscle repair (Wagatsu ma et al. 2011), implying the importance of mitochondrial biogenesis and dynamics during satellite cell differentiation. In aged satellite cells, especially cells obtained from TRI, we have found an altered expression of genes relevant to mitochondrial bi ogenesis, which could at least in part explain their diminished differentiation capacity. Surprisingly, mtDNA copy number, as a marker for mitochondria density, was increased in cells obtained from aged GLU and TRI. An increased in mtDNA copy number has b een reported in various cells in response to aging or oxidative stress ( Gadaleta et al. 1992; Barrientos et al. 1997; Hsin Chen et

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130 al. 2000), and this was explained as a compensatory mechanism for accumulating mtDNA mutations and defects in the OXPHOS system. In our work, lower expression of genes encoding for subunits of OXPHOS complexes in aged cells suggested defects in OXPHOS, which might have induced the increase in mtDNA copy number. However, at present we are not able to distinguish whether the l ower transcript level of mitochondrial genes is an inducer or a consequence of the elevated mtDNA copy number, since the aged cells might have poor transcription efficiency even with higher template availability. It is also noteworthy that the age related elevated mtDNA copy number was not paralleled by increases in CS content, which supports the hypothesized compensatory nature of mtDNA amplification, and speaks against a higher mitochondrial content. Another finding of this work was the accumulation of p 62 in aged satellite cells, indicating an impairment of autophagic degradation. This was further confirmed by elevated LC3II protein level. Involvement of autophagy in cell differentiation has been reported in C2C12 cells ( Sin et al. 2016; Call et al. 20 17 ), where the autophagy process was upregulated soon after cell differentiation was initiated. It is believed that in order to develop a healthy and organized mitochondrial network for the higher energetic demands of myotubes, cells must eliminate the pre existing mitochondria prior to repopulation with newly synthesized ones. Failure of autophagic degradation of existing, possibly impaired, mitochondria during myogenic differentiation may lead to limited energy production for the fusion machinery, and the refore could result in the compromised fusion capacity observed in aged satellite cells obtained from TRI. We hypothesized that the accumulation of p62 and LC3 II was a consequence of decreased expression of LAMP2, which facilitates the fusion of the autop hagosome with the

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131 lysosome (Eskenlinen et al. 2004; Kchl et al. 2006 ). However, we did not detect a significant decrease in LAMP2 gene expression, and the evaluation of protein expression of markers involved in degradation is needed. In conclusion, our current work explored the intrinsic changes in mitochondrial function and quality control in normally aged satellite cells obtained from American Quarter Horses. In the early phase of differentiation, the time period before or upon nasce nt myofibers presented, impaired mitochondrial function was concomitant with, and probably a consequence of, altered mitochondrial quality control. Taken together, these impairments could be a potential cause of compromised myogenic capacity of satellite c ell in vitro Aged satellite cells, especially those derived from TRI started to show alterations in mitochondria quality control mechanisms on day 2 post differentiation. We hypothesize that these alterations might be more pronounced in the late phase of differentiation, which need s to be further studied. Continuing the study of satellite cell biology in healthy aging horses and the pathways that regulate the aging process of the satellite cell itself would help to develop innovative interventions to maint ain muscle mass and function in our aging equine companions.

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132 Figure 4 1 Proliferation rate of satellite cells isolated from skelatlal muscle of American Quarter Horses. Cell number measured by direct count of viable cells in a hemocytometer. Satellite cells were grown for 3, 4, 5, or 6 days in growth media. Data for cell count/well are mean s SE (n = 4 for each group; 3 wells per sample at each time point). Young vs. aged: P < 0.1 P < 0.05

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133 Figure 4 2 Expresion of myogenin in satellite cells isolated from skeletal muscle of American Quarter Horses. Satellite cells isolated from young and aged muscles were differentiated for 3 days, stained for myogenin and counterstained with a nuclear stain (Hoechst). The percentage of myogenin + cells is shown as mean s SE (n = 4 for each group) Open bars represen t young horses; solid bars, aged horses. Young vs. aged: P < 0.1, P < 0.05.

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134 Figure 4 3. Fusion capacity of satellite cells isolated from skeletal muscle of American Quarter Horses. Satellite cells isolated from young and aged muscles were differentiated for 3 days, stained for MyHC and counterstained with a nuclear stain (Hoechst). The fusin capacity is indicated by fusion index that is calculated as t he percentage of nuclei present in MyHC + cells Values are mean s SE ( n = 3 4 for each group) Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.05.

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135 Figure 4 4 The protein expression of citrate synthase in satellite cells isolated from skeletal muscle of American Quarter Horses. A) Representative Western blot image of whole cell lysates at day1 and day 2 postdifferentiation. B C) Quantification of western blot da ta normalized to tubulin Values are mean s SE ( n = 3 4 for each group) Open bars represent young horses; solid bars, aged horses.

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136 Figure 4 5 Mitochondrial DNA copy number in satellite cells isolated from skeletal muscle of American Quarter Horses. Satellite cells isolated from young and aged m uscles were differentiated for 2 days, with a m itochondrial DNA copy number was determined every 24h using quantitative real time PCR of ND1 (A and C ) and COX1 (B and D ), normalized to nuclear DNA copy number actin gene ( ACTB ). Values are mean s SE ( n = 4 for each group) Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.1, P < 0.05.

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137 Figure 4 6 Transcript levels of mtDNA encoded genes in differentiated satellite cells. Gene expression of ND1 (A and D ), COX1 (B and E ), COX2 (C and F ) at D1 DM and D2 DM. Values are mean s SE ( n = 4 for each group) Open bars represent young horses; solid bars, age d horses. Young vs. aged: P < 0.05, ** P < 0.01.

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138 Figure 4 7 Protein expression of Hsp60 in satellite cells during differentiation in vitro Densitometric quantification of Hsp60 protein expression in satellite cells derived from young and aged horses. Representative Western blot images are shown above the graph. Values are mean s SE ( n = 3 4 for each group) Open bars represent young horses; solid bars, aged horses.

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139 Figure 4 8 Transcript level of genes relevant to mitochondrial biogenesis in satellite cells during differentiation in vitro Gene expression of PGC NRF1 and TFAM in differentiated satellite cell at D1 DM (A C) and D2 DM (D F). Results are represented as the fold change compared to young GLU muscle. Values are mean s SE ( n = 4 for each group) Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.1, P < 0.05, ** P < 0.01.

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140 Figure 4 9 P rotein expression of autophagy regulators in satellite cells during differentiation in vitro A) Representative Western blot images of autophagy regulators at day1 and day 2 postdifferentiation. Quantification of p62, LC3I, and LC3II in satellite cells derived from young ( n = 3 4 for both GLU and TRI ) and aged (n = 3 4 for both GLU and TRI ) horses at D1 DM (B D) and D2 DM (E G). Values are mean s SE. Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.1, P < 0.05

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141 Figure 4 9. Continued

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142 Figure 4 10 Gene expression of LC3 in satellite cells during differentiation in vitro Expression of LC3 mRNA in satellite cells isolated from young ( n= 4 for both GLU and TRI ) and aged ( n = 4 for both GLU and TRI ) horses at D1 DM (A) and D2 DM (B). Data are represented as the fold change compared to young GLU muscle. Values are mean s SE. Open bars represent young horses; solid bars, aged horses. Young vs. aged: P < 0.05.

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143 Figure 4 11 Transcript level of the LAMP2 gene in satellite cell isolated from skeletal muscle from American Quarter Horses Expression of LAMP2 mRNA in satellite cells at D1 DM (A) and D2 DM (B) is represented as the fol d change compared to young GLU muscle. Values are mean s SE (n = 3 4 for each group) Open bars represent young horses; solid bars, aged horses.

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144 CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTIONS As the number of aged horses increases aged horse population contributes significantly to the whole equine population. With the willingness of owners to use and work with their older horses, understanding the physiology of equine aging and underlying mechanisms is warranted. Yet, few aging studies have been done in the horse. The work in this dissertation has started to ev aluate differences in skeletal muscle energy metabolism, especially mitochondrial function, between young and old horses in order to better understand the impact s of aging on equine skeletal muscle. Effects of aging on skeletal muscle oxidative capacity was first examined in Chapter 2, we conclude that equine skeletal muscle aging was associated with an increase in percentage of type I and IIA fibers, and a decrease in percentage of type IIX fibers. M itochondrial content and enzymatic activity, as indicated by CS and COX activity respectively, were decreased with age. Compromised mitochondrial function was observed on the mitochondrial level, but not on the muscle level. In Chapter 3, aging related decline in mitochondrial content in equine skeletal muscle was further confirmed and underlying mechanisms responsibl e for compromised mitochondrial function was explored. Decline in expression level of biogenesis bio markers in aged TRI muscle suggest s tha t mitochondrial biogenes is declined with age in equine TRI muscle. In addition, dysregulation of autophagic proteins in TRI muscle indicates that the autophagic flux was impaired in aged TRI muscle. Taken together, decline in mitochondrial biogenesis activ ity and autophagic capacity could be the potential cause of impaired mitochondrial function in aged equine skeletal muscle

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145 After investigat ing the effects of aging on equine skeletal muscle, we asked how muscle stem cells ( namely satellite cells, SCs) were affected by aging in Chapter 4 The ability to regenerate muscle plays a major role in muscle homeostasis after birth and this regenerative capacity is own to muscle SCs Primary culture of SCs under standard conditions in vitro enables us to gain in sight into the intrinsic changes of SC function with age. Our data showed that SC derived from aged horses had decreased proliferative and differentiation capacity compared to those from young horses In line with the decline in SC regenerative capacity, t he mitochondrial OXPHOS seemed to decrease with age as indicated by the lower transcript level of mitochondrial genes in SC from aged horses compared to those from young horses. Moreover, impaired mitochondrial biogenesis and autophagic flux were observed in SCs derived from aged horse. Based on our collective data, we conclude aging was accompanied by compromised mitochondrial function in equine skeletal muscle and satellite cells. However there was no overt mitochondrial dysfunction on whole muscle leve l as we found in Chapter 2. If a transition age exists in the horse, defining it might provide insights about a beneficial time point to apply interventions aiming to delay the onset of overt muscle oxidative dysfunction and decline in physical performance On the other hand, if the horse ages differently from traditional animal models and humans, it will be of interest to charact erize the underlying differences However, it is unlikely that horse ages differently, since we have detected early signs of skel etal muscle aging. Therefore, examining aging effects in even older horses might help to better understand the equine aging process. Moreover, systematically understanding of the underlying mechanisms

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146 responsible for the maintenance of a healthy mitochondr ial population in equine skeletal muscle is a prerequisite to design interventions to prolong health and performance of aging horses. A n even further step will be i f and how interventions that target mitochondrial biology (such as exercise training and resveratrol) affect mitochondrial function in older horses

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147 APPENDIX A CITRATE SYNTHASE ACTIVITY PROTOCOL ** Adapted from Spinazzi et al. ( Spinazzi et al ., 2012) ** To prepare muscle homogenate: 1. Add 10 15 mg powdered (cryopulverized) muscle to each microvial, recording exact tissue weight 2. Add the volume of sucrose homogenization buffer with detergent required to obtain a 40 fold dilution a. Sucrose homogenization buffer: 20 mM Tris, 40 mM KCl, 2 mM EGTA, 250 mM sucrose, pH 7.4 b. Dilute 1 part 5% detergent (n D maltoside; Sigma D4641) to 100 parts sucrose buffer c. with detergent 3. Sonicate (F60 Sonic Dismembrator, Fisher Scientific, Waltham, MA) each tube by pushing the button on the top of the probe for 1sec, 5 10 times, with the strength between 3 5. a. Keep on ice while sonicating b. Clean sonicator probe in between samples by rinsing with dH 2 O 4. Centrifuge microvials for 3 min at 10,000 x g at 4 C 5. Collect supernatant (40 fold dilution) 6. Dilute to 80 fold (1part 40 fold+1part sucrose buffer without detergent (2a)) 7. Can stor e 40 fold and 80 fold homogenates at 80C until analysis 8. Use 80 fold dilution for assay To perform assay: 1. Prepare 200 mM Tris ( pH 8.0) with Triton X (0.2% (vol/vol)) a. Dissolve 1.21 g of Tris in 40 mL distilled H 2 O, adjust to pH 8.0 with HCl, add 0.1 mL of Triton X and adjust the volume to 50 mL b. C an be stored at 4C for up to 2 mo 2 Prepare 1 0 mM Acetyl CoA a. Dissolve 100 mg Acetyl CoA (CHEM IMPEX INT'L INC #00583) in 11 .35 mL distilled H 2 O b. Can be stored at

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148 c. Once thawed, use same day 3 Prepare 1 mM DTNB a. Dissolve 7.9 mg DTNB (Sigma # D8130) in 20 mL of 100 mM Tris (pH 8.0) b. Prepare fresh daily 4 Turn on microplate reader, check protocol (CS Spinazzi in Gen5 in Wohlgemuth lab; be sure pathlength correction is INACTIVATED), and preheat to 37C 5 Prepare 10 mM oxaloacetic acid (OAA) a. Dissolve 6.6 mg OAA (Sigma # O4126) in 5 mL distilled H 2 O b. Prepare fresh daily 6 Prepare reaction mix a. Per well: i. 2 O ii. X (0.2% (vol/vol) from step 1 ) iii. tep 3 ) iv. Mm from step 2 ) 7 8 Add 23 ) to each well 9 Read baseline activity at 412 nm for 3 min 10 Start reaction by addin ) to each well using multichannel pipette one column at a time 11 Monitor increase in absorbance at 412 nm for 3 min To get pathlength: When the plate is completed, save data and close off the current protocol. 1. Re open the same protocol you just used (CS Spin azzi in Gen5 in Wohlgemuth lab) but be sure path length correction is ACTIVATED 2. Read the pla te until you get the pathlength To calculate activity: 1. Calculate slope (change in absorbance over time) for baseline reading and activity read ing for each sample 2. Subtract baseline slope from activity slope

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149 3. CS activity (nmol/min/mg protein) = t otal volume (0.250) (cm ) x s ample volume (0.004) x total protein in sample (mg/mL)

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150 APPENDIX B CYTOCHROME C OXIDASE ACTIVITY PROTOCOL **Adapted from Spinazzi et al., (Spinazzi et al ., 2012)** To prepare muscle homogenate: 1. Add 10 15 mg powdered (cryopulverized) muscle to each microvial, recording exact tissue weight 2. Add the volume of sucrose homogenization buffer with detergent required to obtain a 40 fold dilution a. Sucrose homogenization buffer: 20 mM Tris, 40 mM KCl, 2 mM EGTA, 250 mM sucrose, pH 7.4 b. Dilute 1 part 5% detergent (n D maltoside; Sigma D4641) to 100 parts sucrose buffer c. with de tergent 3. Sonicate (F60 Sonic Dismembrator, Fisher Scientific, Waltham, MA) each tube by pushing the button on the top of the probe for 1sec, 5 10 times, with the strength between 3 5. a. Keep on ice while sonicating b. Clean sonicator pr obe in between samples by rinsing with dH 2 O 4. Centrifuge microvials for 3 min at 10,000 x g at 4C 5. Collect supernatant (40 fold dilution) 6. Dilute to 80 fold (1part 40 fold+1part sucrose buffer without detergent (2a)) 7. Can store 40 fold and 80 fold homogenates at 80C until analysis 8. Use 80 fold dilution for assay To perform assay: 1. Turn on microplate reader, check protocol (COX Spinazzi in Gen5 in Wohlgemuth lab; be sure pathlength correction is INACTIVATED ), and preheat to 37C 2. Prepare 100 mM pota ssium phosphate buffer ( pH 7.0) a. Titrate 100 mM potassium phosphate dibasic with 100 mM potassium phosphate monobasic up to a pH of 7.0 b. C an be store d at 4C for up to 2 mo

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151 3. Prepare 1mM reduced cytochrome c a. To make 1mM oxidized cytochrome c by dissolving 12.5 mg oxidized cytochrome c (Sigma # C7752) in 1 mL of 20 mM potassium phosphate buffer (20 mM) b Reduce cytochrome c solution with a few grains of dithionite (pipette tip) just before use c. Vortex thoroughly (will change color from brown to orange pink) 4 Prepare reaction mix a. Per well: i. ii. from step 2 ) iii. c (from step 3 ) 5 Add 270 ) to each well of a background plate a. Read baseline activity of background plate at 550 nm for 10 min b. Keep background plate in thermomixer at 37C after baseline read 6 a. using multichannel pipette one column at a time b. Monitor decrease in absorba nce at 550 nm for 3 min To get pathlength: When the plate is completed, save data and close off the current protocol. 1. Re open the same protocol you just used (COX Spinazzi in Gen5 in Wohlgemuth lab) but be sure pat hlength correction is ACTIVATED 2. Read the plate until you get the pathlength To calculate activity: 1. Calculate slope (change in absorbance over time) for baseline reading and activity reading for each sample 2. Subtract baseline slope from activity slope 3. COX activity (nmol/min/mg protein) = Absorbance/min x1000) x t otal volume (0.250) (cm ) x s ample volume (0.005) x total protein in sample (mg/mL)

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152 APPENDIX C 3 OH ACYL COA DEHYDROGENASE ACTIVITY PROTOCOL **Adapted from Fong & Schulz (Fong & Schulz, 1978)** To prepare muscle homogenate: 1. Add 10 15 mg powdered (cryopulverized) muscle to each microvial, recording exact tissue weight 2. Add the volume of sucrose homogenization buffer with detergent required to obtain a 40 fold dilution a. Sucrose homogenization buffer: 20 mM Tris, 40 mM KCl, 2 mM EGTA, 250 mM sucrose, pH 7.4 b. Dilute 1 part 5% detergent (n D maltoside; Sigma D4641) to 100 parts sucrose buffer with detergent 3. Sonicate (F60 Sonic Dismembrator, Fisher Scientific, Waltham, MA) each tube by pushing the button on the top of the probe for 1sec, 5 10 times, with the strength between 3 5. a. Keep on ice while sonicating b. Clean sonicator probe in between samples by rinsing with dH2O 4. Centrifuge microvials for 3 min at 10,000 x g a t 4C 5. Collect supernatant (40 fold dilution) 6. Dilute to 80 fold (1part 40 fold+1part sucrose buffer without detergent (2a)) 7. Can store 40 fold and 80 fold homogenates at 80C until analysis 8. Use 80 fold dilution for assay To perform assay: 1. Turn on microplate reader, check protocol ( 3 HADH in Gen5 in Wohlgemuth lab; be sure pathlength correction is INACTIVATED), and preheat to 37C before use 2. Prepare 5 0 mM potassium phosphate buffer (pH 7 .0 ) with 0. 06% (vol/vol) Triton X 100 a. Titrate 5 0 mM pot assium phosphate dibasic with 5 0 mM potassium phosphate monobasic up to a pH of 7.0 b. Add Triton X

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153 c. Can be store d at 4C for up to 2 mo 3. Prepare 0.9 mM Acetoacetyl coenzyme A sodium salt hydrate (SAAC) a. Dissolve 1.533 mg SAAC (Sigma # A1625) in 2 mL of 50 mM potassium phosphate buffer (pH 7.0) b. Can be stored at 4C for up to 2 mo Nicotinamide Adenine Dinuc leotide, reduced disodium salt hydrate (NADH) a. Dissolve 5.108 mg NADH (Sigma # N8129) in 1 mL of 50 mM potassium phosphate buffer (pH 7.0) b. Prepare fresh just before use 5 Prepare reaction mix a. Per well: i. 100 (from step 2) ii. ( 7.2 mM from step 4) ll 8. Read baseline activity at 340 nm for 2 min multichannel pipette one column at a time 10. Monitor decrease in absorbance at 340 nm for 7 min To get pathlength: When the plate is completed, save data and close off the current protocol. 1. Re open the same protocol you just used ( HADH in Gen5 in Wohlgemuth lab) but be sure pathlength correction is ACTIVATED 2. Read the plate until you get the pathlength To calculate activity: 1. Calculate slope (change in absorbance over time) for baseline reading and activity reading for each sample 2. Subtract baseline slope from activity slope

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154 3. NADH activity ( n mol/min/mg protein) = otal volume (0.275 ) ) x pathlength (cm) x sample volume (0.010 ) x total protein in sample (mg/mL)

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155 APPENDIX D MYOSIN HEAVY CHAIN ISOFORMS IDENTIFICATION PROTOCOL ** Adapted from Talmadge & Roy ( Talmadge & Roy, 1993)** Part I G el casting column except TEMED and 10% AP, then vortex the solution and degas. 2. Add TEMED and 10% AP to the solution made in step1 and pour to the gel cassette to the mark (a black line draw n 1cm below the comb teeth). **pour the solution smoothly to prevent air bubbles** 3. Immediately ov erlay the solution with water. **add water slowly and evenly to prevent mixing** 4. Allow the gel to polymerize for 40 min to 1h 5. When the gel is solid enough, remove the water and rinse the gel surface completely with d water. Table D 1. SDS PAGE gel mixture Stock solution Separating gel (mL) Stacking gel (mL) 100% glycerol 3.0000 1.5000 30% Acrylamide bis (50:1) 2.6670 0.6650 Tris HCL 1.5M (pH 8.8) 1.3300 Tris HCL 0.5M (pH 6.8) 0.7000 1M glycine 1.0000 100 mM EDTA (pH 7.0) 0.2000 10% (w/v) SDS 0.4000 0.2000 Distilled H2O 1.4950 1.6810 TEMED 0.0050 0.0025 10% Ammonium persulfate (daily fresh) 0.1000 0.0500 except TEMED and 10% AP, then vortex the solution and degas. 7. Before casting the stacking gel, insert a piece of filter paper to dry the area between the two glass plates above the separating gel. **do not tough the surface of the separating gel ** 8. Add TEMED and 10% AP to the solution made in step 6 and pour to the gel cassette until the topof the short glass plate. **pour the solution smoothly to prevent air bubbles**

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156 9. Insert the desired comb, making sure no air bubbles. 10. Allow the stacking gel to polymerize for 45 min. 11. Gently remove the comb and rinse the wells thoroughly with d water or running buffer Part II E lectrophoresis 1. Assemble the Mini Protean 3 (follow the gel casting manual) 2. buffer (100 mM Tris + 150 mM glycine + 0.1% SDS) chamber until merging the wells 3. Take the supernatant from 20C, and mix with Laemmli buffer 4. Heat the sample mixture for 3 min at 95 C, then leave at RT unti l loading 5. Lo ad samples to wells 6 (50 mM Tris + 75 mM glycine + 0.05% SDS) Tank (outer) 7. Run electrophoresis for 18 h at 4C ( run the gel at 120 V for the first 2 h, then at 100 V for the rest 16 h) 8. Remove the gel from electrophoresis chambers and start the Coomassie blue staining Part III Coomassie blue staining 1. After electrophoresis place gel in a plastic container and overlay with Gel Fix Solution so that gel floats easily. Agitate on an orbital shaker or rocking platform for 0.5 h 2. Remove Gel Fix Solution and verlay gel in at lea st 100 ml of CBB R 250 Solution and agitate on an orbital sh aker or rocking platform 2 h 4. A fter staining, wash the gels with several changes of water 5. Place gel in at least 100 ml of Detain Solution and agitate on an orbital shaker or rocking platform until resolved blue bands and a clear backgroun d appear (~ 2 h ). ** Destaining must be monitored visually and adjusted accordingly ** 6. Place gel in Gel Storage Solution

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157 7. Photograph the gel or analyze the gel spectrophotometrically 8. Buffer recipes for staining a. GEL FIX SOLUTION : i. 50% Methanol, 40% dH2O, 10% Glacial acetic acid ii. Example: Add 250 mL methanol to 200 mL dH 2 O. Slowly add 50 mL glacial acetic acid iii. Can be s tore d at RT for several mo b. CBB R 250 SOLUTION: i. 0.1% CBB R 250, 40% Methanol, 50% dH2O,10% Glacial acetic acid ii. Example: Dissolve CBB R 250 in 200 mL Methanol. Add 250 mL dH 2 O then 50 mL glacial acetic acid iii. Can be stored at RT in a dark bottle for several mo c. DESTAIN SOLUTION i 10% Methanol 83% dH 2 O 7% Glacial acetic acid ii. Example: Add 50 mL methanol to 415 mL dH 2 O. Slowly add 35 mL g lacial acetic acid iii. Can be stored at RT for several mo d. GEL STORAGE SOLUTION i. 95% dH 2 O 5% Glacial acetic acid ii. Example: Add 25 mL glacial acetic acid to 475 mL dH 2 O iii. Can be stored at RT for several mo

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158 APPENDIX E PERMEABILIZED FIBERS PROTOCOL **Adapted from E. Gnaiger/Oroboros** Part I Muscle fiber preparation 1. Prepare BIOPS buffer according to Table E 1 Table E 1. BIOPS buffer, total volume = 1 L Compounds in BIOPS Final conc. Stock solution Addition to 1 L final Source and product code CaK 2 EGTA 2.77 mM 100 mM 27.7 mL K 2 EGTA 7.23 mM 100 mM 72.3 mL Na 2 ATP 5.77 mM 3.141 g Sigma A2383 MgCl 2 6H 2 O 6.56 mM 1.334 g Scharlau MA0036 Taurine 20 mM 2.502 g Sigma T0625 Na 2 Phosphocreatine 15 mM 4.097 g Sigma P7936 Imidazole 20 mM 1.362 g Fluka 56750 DTT 0.5 mM 0.077 g Sigma D0632 MES hydrate 50 mM 9.760 g Sigma M8250 2. P repare MiR05 b uffer according to Table E 2 Table E 2 MiR05 buffer, total volume = 1 L Compounds in MiR05 buffer Final conc. Addition to 1 L final Source and product code EGTA 0.5 mM 0.190 g Sigma E4378 MgCl 2 6H 2 O 3 mM 0.610 g Scharlau MA0036 Lactobionic acid 60 mM 120 Ml of 0.5 M K lactobionate stock Aldrich 153516 Taurine 20 mM 2.502 g Sigma T0625 KH 2 PO 4 10 mM 1.361 g Merck 104873 HEPES 20 mM 4.770 g Sigma H7523 D Sucrose 110 mM 37.650 g Sigma 84097 BSA, essenrially fatty acid free 1 g/L 1 g Sigma A6003 3 Teasing of Muscle Fibers The preparation (teasing) of muscle fibers for respiration measurements should not exceed 30 min and should proceed on ice a. Dissect (25 mg of) muscle tissue or receive muscle biopsy specimen and place immediately in ice cold BIOPS Buffer Muscle sample CANNOT dry out For muscle fiber preparation, place muscle sample in 100 cm culture dish on ice, and

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159 position under dissecting scope b. Trim muscle of connective tissue and cut muscle longitudinally into smaller layers (ideally there will be fiber monolay ers, but this might not be feasible with biopsy samples) c. Under a dissecting microscope, and using a pair of needle tipped forceps (Dumont #5), separate fibers from one another to maximize the surface area of the fibers that saponin can access, leaving only small regions of contact. The passive motion of the opening forceps can be used to separate the fibers. Teasing should not take no longer than 10 15 min 4 Muscle fiber permeabilization a. To permeabilize the prepared muscle fibers, place each fiber bundle in ice cold BIOPS buffer and incubate on rotator for 30 min at 4C (inside fridge). Make sure that the muscle pieces are moving when tube is on the rotator i. A dd saponin stock solution (5 mg/mL) into vial with BIOPS buffer right before BIOPS buffer b. To wash permeabilized fiber bundles, place bundle as best and complete as possible to a new tube containing ice cold MiR05 buffer for 10 min on rotator at 4C to remove saponin and an y extramitochondrial components Part II Oxygen consumption measurement 1. Adding the tissue into the chamber and getting started a. When the O2k is prepared, weigh 2 3 mg of fibers by carefully blotting fiber bundles on Kimwipe and putting on weigh paper in Analytical Balance. Make note of the weight b. Transport fibers to O2k by carrying weigh boat or weighing paper with fibers to O2k c. A dd the weighed fibers to the chamber containing MiR05 containing 20 mM creatine d. measurement e. As soon as the fibers are in, turn the stirrers back on and insert the stoppers, (semi open) (using s pacer) f. A dd 10 20 mL pure O 2 gas through the stopper using special syringe and length adapted needle g. Watch the O 2 concentration rise (blue line) and insert the stopper completely (closing the chamber) when O 2 concentration will continue i. Make sure that the chamber does not contain any air bubbles since this will disturb and falsify the O 2 flux signal ii. Do not let the O 2 concentration fall below 220 open the chamber (use spacer) and, in case of high O2 consumption, reinject O 2 gas until O 2

PAGE 160

160 2. Oxygen Flux measurements Titration Protocol a. Some commonly used substrates, inhibitors an uncouplers are listed in Table E 3 Table E 3 Commonly used SUIT chemicals Substrate Stock conc. in syringe Final conc. in 2 mL Titration ( ) Storage Pyruvate 2 M (in H 2 O) 5 mM 5 Make fresh Malate 0.8 M (in H 2 O) 2 mM 5 Stored at 20 C Glutamate 2 M (in H 2 O) 10 mM 10 Stored at 20 C Succinate 1 M (in H 2 O) 10 mM 20 Stored at 20 C Cyt c 4 mM (in H 2 O) 5 Stored at 20 C ADP 0.5 M (in H 2 O) 1 5 mM (2 mM) 4 20 (8) Stored at 80 C Oligomycin 5 mM (EtOH) 1 Stored at 20 C FCCP 1 mM (EtOH) Stored at 20 C Antimycin A 5 mM (EtOH) 1 Stored at 20 C b Titration protocol (SUIT protocol) Table E 4 SUIT protocol used in C hapter 2 Titration substrates (final conc. in 2 mL; titration vol) Response Glutamate and Malate (10/2 mM; L; LEAK; Small increase and stabilization. Wait for a 5 10 min stable flux ADP (2.5 mM; P CI ; Rapid increase. Wait 5 10 min after the signal stabilized. P CI+II ; Elevation above P CI Cytochrome c P CI+II c ; No increase or small elevation of flux ADP and Succinate (1.25/5 mM; whether ADP or Succinate was limiting titrations) E; ETS; Elevation above P (excess capacity) or just to P CI+II ROX, residual (non mitochondrial) respiration. Flux will decrease below LEAK level

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161 Part III Post measurement device c leaning 1. dH 2 O washes (3x) a. T ake out sto ppers and rinse stopper and capillary with water several times (5x) b. S iphon off the media from chamber c. F ill with water (stirrer on) and siphon off d. R epeat 2 more times 2. EtOH washes (3x w/70% EtOH + 1x w/100% EtOH) a. R inse stopper from outside and a capillary with 70% EtOH several times (3x) b. S iphon off the water from the chamber and fill with 70% EtOH, insert stopper until receptacle fills with the repeat EtOH; incubate for 5 min while stirring c. R epeat 2 more times d. S iphon off the 70% EtOH and replace with 100% EtOH, insert stopper until receptacle fills with EtOH; incubate for 15 min while stirring e. P roceed to (4) for storage or to (3) for continuation of experiments 3. In preparation of the next experiment: Make sure the stirrer is rotating. Remove the stopper, rinse the surface and cannula of the stopper with dH 2 O. Place the stopper clean and securely (in the tube with distilled water). Rinse the chamber with disti lled water three times. Then proceed with the next experimental run. 4. For storage: Fill the chamber with 70% EtOH. Insert the stopper loosely and fill 70% EtOH up to the rim of the receptacle. Place the Cover onto the stopper to minimize evaporation and leakage of EtOH. For overnight storage and chemical sterilization keep EtOH in the chamber and switch off the O2k. You can use this method for storage up to several months, with the POS in place, ready for use.

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195 BIOGRAPHICAL SKETCH She finished her high school at her hometown, where she was an honor student and received scholarships several times. In 2005, she attended the Shandong Agricultural University with a major in animal science. In the four years she spent in college, she received continued Academic Excellence Scholarship and graduated in 2009 Following that, Chengcheng pursued a Ma s ter of Science degree in animal nutrition and feed science program, where she started to get interested in the cellular and molecular biology. In 2012, she graduated with an Outstanding Graduate Student Award. Afte r taking a year off from school, Chengcheng was admitted at the University of Florida under the Doctor of Philosophy in Animal Molecular & Cellular Biology program. During her PhD life, Chengcheng received Graduate Assistanship fellowship. She was also awa rded Outstanding Academic Achievement and Susan Meg Weinstein Animal Sciences Graduate Scholarship and she completed her PhD in the summer of 2017 Upon completion of her PhD degree, Chengcheng plans to continue biology research as a post doctoral scholar in Unite d States and explore more in the molecular and cellular biology related fields.