Physiological and Pathophysiological Regulation of the Ryanodine Receptor in Skeletal Muscle

Home , AKAP6, KCNE1, MIP (gene), Ryanodine receptor 2

Physiological and pathophysiological regulation of the ryanodine receptor in skeletal muscle

Alisa Umanskaya

Submitted in partial fulfillment of the

requirements for the degree of

Doctor of Philosophy

in the Graduate School of Arts and Sciences

COLUMBIA UNIVERSITY

2015

Abstract

Physiological and pathophysiological regulation of ryanodine receptor

in skeletal muscle

Alisa Umanskaya

Ryanodine receptor calcium release channels are essential for skeletal muscle contraction, as they mediate the release of calcium ions from intracellular stores into the cytosol. The data presented in this dissertation demonstrate the evolutionarily conserved mechanisms of skeletal muscle ryanodine receptor regulation in the physiological and pathophysiological states.

Adrenergic stimulation causes increased skeletal muscle force, however, despite the well- established role of this physiological response, the molecular mechanism is not known.

Here we present a mechanism whereby phosphorylation of a single amino acid on the ryanodine receptor is a key signal in the physiological stress-induced inotropic response in mouse skeletal muscle. Therefore acute post-translational modifications of ryanodine receptor channels are important for healthy muscle contraction. Conversely, chronic stress-induced post-translational modifications result in poorly functioning murine ryanodine receptor channels that contribute to skeletal muscle dysfunction in age- dependent skeletal muscle weakness and Muscular Dystrophies.

Finally, we present data that demonstrates striking evolutionary conservation in ryanodine receptor regulation in the physiological and pathophysiological states between mice and C. elegans. This work has broad implications for understanding the underlying mechanisms of skeletal muscle contraction and important disorders that affect human health. Furthermore, this works presents ryanodine receptor channels as a viable therapeutic target for age-related skeletal muscle weakness, Muscular Dystrophies, and also implicates C. elegans as a potential model system in which to test future therapeutic targets.

Table of Contents

List of Figures……………………………………………………………………………..ii

Chapter 1: Introduction…………………………………………………………………… 1 v Mammalian excitation-contraction coupling……………………………………... 1 v Ryanodine receptors……………………………………………………………… 6 v Ryanodine receptor regulation by post-translational modifications and regulatory proteins………………………………………………………………....………... 11 v Ryanodine receptor redox sensing………………………….…………………… 16 v Ryanodine receptor-related skeletal muscle pathophysiology………….……….. 21 v Excitation-contraction coupling in C. elegans……….…………….……………. 23 v Redox balance/imbalance in aging C. elegans………………………….………..28 v Rationale and aims………………………………………………………………. 30

Chapter 2: Post-translational modification of the ryanodine receptor type 1 underlies skeletal muscle weakness in the physiological and pathophysiological state…………… 38 v Phosphorylation of the ryanodine receptor by protein kinase A is a molecular mechanism underlying fast twitch skeletal muscle inotropy……………………. 38 v Post-translational modifications of the ryanodine receptor mechanistically underlie skeletal muscle weakness in Limb-girdle muscular dystrophy...……… 45

Chapter 3: Skeletal Muscle Aging: An acquired form of Muscular Dystrophy...………. 53 v Ryanodine receptor oxidation causes muscle weakness in aging. ...……………. 53 v Genetically enhancing mitochondrial anti-oxidant activity improves muscle function in aging.……………………………...... ……………………………… 77

Chapter 4: C. elegans: A model for the physiological study of muscle function in aging…………………………………………………………………………………… 104 v The role of the UNC-68/intracellular calcium release channel in age- dependent loss of muscle function in C. elegans………………………. 104

Discussion…………………………...... ………………………………………………. 120

References………………………………………………………………………………132

List of figures

Figure 1: Cardiac vs. skeletal muscle E-C coupling.……………………………….…… 33

Figure 2: Physiological vs. pathophysiological skeletal muscle E-C coupling…………. 34

Figure 3: Oxidative systems in mammalian skeletal muscle and C. elegans…………… 35

Figure 4: Schematic representation of the dystrophin glycoprotein complex (DGC)…... 36

Figure 5: Sequence identity between mammalian and C. elegans ryanodine receptors and immunophillins...…………………………………….……………………..………….... 37

Figure 6: β-adrenergic pathway………………………………….……………….……... 53

Figure 7: RyR1-S2844A muscle exhibits similar force to WT muscle…………………. 54

Figure 8: β-adrenergic inotropy in skeletal muscle is dependent on phosphorylation of serine 2844 in RyR1.………………………………….……………….……...... ………. 55

Figure 9: Blocking phosphorylation of RyR1 at S2844 inhibits the β-adrenergic effect on Ca2+ transients in FDB muscle fibers. ……….…….……….…….……….…….………. 56

Figure 10: Abolished phosphorylation of RyR1 following β-adrenergic stimulation and reduced in vivo force production in S2844A mice..…….……….…….………………… 57

Figure 11: RyR1 in β-sarcoglycan deficient muscle is cysteine-nitrosylated, oxidized and depleted of calstabin1.…….……….…….…………….……….…….…………………. 58

Figure 12: β-sarcoglycan deficient muscle displays defective SR Ca2+ release that is restored by S107 treatment..…….……….…….……….…….……………….…..…….. 59

Figure 13: S107 treatment increases muscle force in β-sarcoglycan deficient mice……..60

Figure 14: S107 treatment increases voluntary exercise in β-sarcoglycan deficient mice..…….…………….………………….………………….………………….……….61

Figure 15: EDL muscles from β-sarcoglycan deficient mice exhibit abnormal mitochondrial morphology.……………....………………….…………………………...62

Figure 16: Impaired force production and reduced SR Ca2+ release in EDL muscle from aged mice.……………....………………….…………….………………………... 90

Figure 17: RyR1 from aged muscle is cysteine-nitrosylated, oxidized and depleted of calstabin1..………………....………………….…………....………………….………... 91

Figure 18: Pharmacological dissociation of RyR1 and calstabin1 causes increased mitochondrial Ca2+……....………………….…………....………………….…………... 92

Figure 19: Pharmacological dissociation of RyR1 and calstabin1 causes reduced mitochondrial membrane potential.……....………………….……….……….………… 93

Figure 20: Pharmacological dissociation of RyR1 and calstabin1 causes increased mitochondrial ROS and RNS production.….……….…………….……….……………. 94

Figure 21: Improved exercise capacity and muscle specific force following S107 treatment of aged mice…………………..….……….…………….……….……………. 95

Figure 22: Increased SR Ca2+ release following S107 treatment of aged mice..………... 96

Figure 23: Model of RyR1-mediated SR Ca2+ leak and mitochondrial dysfunction in aging skeletal muscle.……………..….……….……………..……………..….………... 97

Figure 24: Improved exercise capacity in aged MCat mice.……………..……………... 98

Figure 25: Preserved skeletal muscle function in aged MCat mice.…..……..…………. 99

Figure 26: Improved tetanic Ca2+ in skeletal muscle from aged MCat mice…..………. 100

Figure 27: SERCA1 activity in MCat mice is similar to WT..……………..………….. 101

Figure 28: Reduced SR Ca2+ spark activity and increased SR Ca2+ load in muscle from aged MCat mice...……………..…………..………..……………..………..………….. 101

Figure 29: Skeletal muscle RyR1 isolated from aged MCat mice is biochemically remodeled and exhibits reduced single-channel open probability (Po)…………………102

Figure 30: UNC-68 is comprised of a macromolecular complex……………………… 113

Figure 31: Age-dependent reduction in Ca2+ transients is accelerated in fkb-2 relative to WT………………………………………………………………………………………114

Figure 32: Age-dependent UNC-68 remodeling is accelerated in fkb-2 relative to WT………………………………………………………………………………………115

Figure 33: Depleting FKB-2 from UNC-68 causes UNC-68 oxidation.…………….…116

Figure 34: FKB-2 depletion from UNC-68 causes defective Ca2+ handling………...…117

Figure 35: UNC-68 oxidation causes defective Ca2+ handling…………….………...…118

iii

Figure 36: Treating aged nematodes with the RyR-stabilizing drug, S107, increases body wall muscle Ca2+ transients in aged C. elegans.…………….………...... …119

Supplementary Figure 1: Confirmation that FKB-2 is part of the UNC-68 macromolecular complex…………………………………………………...... …………...121

Acknowledgements

I consider myself lucky to have found my way into Dr. Marks’ laboratory and to have had the opportunity to be mentored by him. His passion for scientific inquiry is infectious and I hope to embody that enthusiasm and wisdom in my future endeavors.

Furthermore, I would like to thank my lab mates, past and present, for supporting me technically and mentally with openness and humor throughout my years in the Marks laboratory. It has been a true pleasure to learn from and work with so many true experts and I am truly thankful for all of the guidance and help that they have provided throughout the years. I would especially like to thank Steve Reiken, who in his own way, takes care of the laboratory and the people in it.

The post-doctoral scientists and graduate students in the laboratory of Dr. Martin Chalfie were also instrumental in teaching me the basic methodologies involved in the C. elegans component of my dissertation. Dr. Irini Topalidou, in addition to being a wonderful scientific resource throughout my training, has also become a dear friend.

I would like to thank the members of my committee, Drs. Henry Colecraft, Iva Greenwald, Monica Driscoll, and Alain Lacampagne. Dr. Colecraft was the Director of Graduate Studies throughout most of my time in the Physiology and Cellular Biophysics Department and he has been a kind source of encouragement. Both, Drs. Greenwald and Driscoll have welcomed me into their laboratories and have taken the time to meet with me and help guide my project. I have had the pleasure to briefly collaborate with Dr. Lacampagne, and our interactions have added a level of depth and scientific rigor to my undertakings in the laboratory. Each member of my committee has been instrumental to the development of my graduate training.

Finally, I have had a plethora of excellent scientific mentors in my past. Special thanks to Drs. Gustavo Macintosh at Iowa State University and Carl Nathan at Weill Cornell Medical College, both of whom mentored me at a young age and were instrumental in the development of my interest and commitment to the pursuit of a doctoral degree. Dr. Luiz Pedro Sorio de Carvalho, who I worked with when he was a post-doctoral scientist in Dr. Nathan’s laboratory, was also imperative in my development, and has been a constant source of support and encouragement throughout my academic studies.

To my parents, Regina Umanskaya and Felix Umansky with love

Introduction

The elucidation of the sequence of events during excitation-contraction (E-C) coupling was one of the great advances in physiology of the twentieth century1. However, many aspects of both physiological E-C coupling and the pathophysiological state remain unknown. The Global Burden of Disease Study of 2010 estimated that 1.7 billion people worldwide are affected by musculoskeletal disorders. Among the almost 300 diseases and injuries evaluated in the study, musculoskeletal disorders were ranked as the second greatest cause of disability2,3. Thus the study of the mechanisms underlying musculoskeletal disease is vitally important and may harbor implications for novel therapeutic interventions.

Excitation-contraction coupling

Mammalian E-C coupling is a complex process that couples electrical stimulation with the molecular events leading to muscle contraction. Central to this process, is the release of intracellular Ca2+ into the cytosol and the subsequent increase in cytosolic free Ca2+.

Ca2+ is a common second messenger that regulates many cellular processes, such as contraction, secretion, synaptic transmission, fertilization, nuclear pore regulation, and transcription4,5. The large difference in [Ca2+] between the cytosol (~100 nM in most cells) and the extracellular space/intracellular Ca2+ stores (1-3 mM in most cells), along with the slow cytoplasmic Ca2+ diffusion rate (~15 µm2 s−16), account for the ability of the cell to efficiently elevate local cytosolic Ca2+ levels, which is necessary to trigger important physiological processes.

The first methods used to study Ca2+ transport across cell membranes monitored radioisotope (45Ca) influx/efflux between two defined compartments7. From these initial studies, it became clear that Ca2+ was sequestered into intracellular organelles and that the cytosol represented an intermediary compartment with variable Ca2+ buffering properties. The sarcoplasmic reticulum (SR) was determined to be the intracellular Ca2+ store in striated muscles (endoplasmic reticulum in most other cells; ER) by examining

Ca2+ influx/efflux in chemically or mechanically skinned fibers5. There are two families of Ca2+ release channels that reside on the SR/ER: the ryanodine receptors (RyRs) and inositol 1,4,5-trisphosphate receptors (IP3Rs). The RyR and IP3R proteins share significant sequence identity, which is most notable in their respective pore regions8,9.

E-C coupling is vital to muscle force production, and the sequence of events is similar when comparing skeletal and cardiac muscle, with certain key differences that will be discussed throughout this chapter. E-C coupling in the skeletal muscle begins with depolarization of the plasma membrane via acetylcholine-mediated propagation of an action potential initiated at the neuromuscular junction along the surface membrane

(sarcolemma) of a muscle fiber and into the specialized invaginations of the sarcolemma known as transverse tubules (t-tubules). Membrane depolarization triggers a conformational change of the α1s subunit of voltage-gated dihydropyridine receptors

2+ (DHPRs, L-type Ca channels Cav1.1 and Cav1.2 in skeletal and cardiac muscle, respectively) located on the t-tubules. DHPR activation results in the coordinated release of Ca2+ from the SR through many ryanodine receptor channels (RyR1 and RyR2 in skeletal and cardiac muscle, respectively), located in the adjacent terminal cisternae of

the SR. This depolarization-induced activation of RyR1 leads to a rapid rise of about ten- fold in intracellular free [Ca2+] (from ~100 nM to ~1 µM; Figure 1A).

This physiological increase in intracellular free Ca2+ serves both to facilitate muscle force production and also to regulate the molecular components of E-C coupling. Ca2+ binding

2+ 2+ to RyR1 Ca binding sites increases the open probability (Po) at low cytoplasmic [Ca ]

(~0.3-10 µM) and inhibits the channel at high cytosolic [Ca2+] (millimolar range). This

2+ highly calibrated Ca sensitivity ensures that ryanodine receptor channels have low Po at resting cytosolic [Ca2+] and may also help prevent sustained activation of ryanodine receptor channels at high cytosolic Ca2+ concentrations, leading to appropriate and timely muscle relaxation5.

Upon release from intracellular stores, the free Ca2+ binds to the regulatory protein troponin C on the actin filaments. This causes a conformational change, exposing binding sites for myosin on the actin filaments. Consequently, myosin forms ATP-dependent crossbridges with actin, which slide past each other, shortening the sarcomere and resulting in muscle contraction and force production. The Ca2+ reuptake pump, sarcoplasmic/endoplasmic reticulum calcium ATPase (SERCA1a and SERCA2a in skeletal and cardiac muscle, respectively), pumps Ca2+ back into the SR, lowering the cytosolic [Ca2 +] to baseline levels of ~100 nM, thereby causing relaxation. Thus in skeletal muscle, cellular Ca2+ is cycled between the SR and cytoplasm10,11 (Figure 1A).

2+ In contrast to Cav1.2 in cardiomyocytes, skeletal muscle Cav1.1 do not conduct Ca upon membrane depolarization; instead, voltage-dependent coupling of Cav1.1 activation to muscle contraction occurs through the physical interaction of the Cav1.1 intracellular II-

III loop with RyR112,13. It was observed as early as the 1970s that skeletal muscle continued to contract in the absence of external Ca2+ ions14,15. The asymmetric charge movement necessary for this depolarization-induced activation of RyR1 is generated by the movement of positively charged residues in the S4 segment of the Cav1.1 α1s subunit in response to a change in the electric field across the membrane. The skeletal muscle membrane geometry is well suited for this process; the term, couplon, describes the functional unit that is composed of terminal expansions of SR found in close proximity to a central transverse tubule element16. In adult skeletal muscle this functional unit is known as a triad, and is composed of terminal expansions of SR on either side of a t- tubule element, however different combinations of SR and t-tubule elements have been reported in thin section electron micrographs, from diads (one SR and one t-tubule element) to heptads (four SR and three t-tubule elements) or greater combinations17.

A distinctive feature of the skeletal muscle transduction mechanism is speed. Signal transmission between Cav1.1 and RyR1 must occur during the very brief (∼2 ms) skeletal muscle action potential, and this is the likely reason for the voltage-sensing role of the voltage gated Ca2+ channel in this tissue (Figure 1B). The mechanisms underlying RyR1- and RyR2-mediated E-C coupling differ in that no direct protein–protein interaction

18 between the Cav1.2 and RyR2 is necessary for E-C coupling in cardiac muscle . Thus the cardiac molecular geometry is different. It is estimated that there is one Cav1.2 for every

5–10 RyR2 channels and the RyR2 and Cav1.2 are not aligned in such a highly ordered

19 fashion as RyR1 and Cav1.1 in skeletal muscle . The long action potential (∼100 ms) in cardiac myocytes permits cardiac E-C coupling to occur via Ca2+-induced Ca2+ release

(CICR; Figure 1B).

Cardiac CICR refers to the activation of RyR2 by extracellular Ca2+ that enters cardiomyocytes through Cav1.2. Upon depolarization of the plasma membrane, Cav1.2 are activated, causing them to open rapidly and allow extracellular Ca2+ to flow down its concentration gradient and into the cytoplasm. This influx occurs down an ~10,000-fold concentration gradient from ~1 mM to ~100 nM Ca2+ in the extra- and intracellular spaces, respectively. This process is sensitive to both the speed and amplitude of the applied Ca2+ trigger20,21 (Figure 1C).

The extracellular Ca2+ entering the cell binds to and activates RyR2 channels, which further triggers SR Ca2+ release. However intracellular Ca2+ release from the SR contributes substantially more to the total increase in cytosolic [Ca2+] than does

2+ 22 extracellular Ca influx through Cav1.2 during CICR . Once released into the cytosol,

Ca2+ binds the contractile machinery, causing cardiomyocyte contraction (systole).

Muscle relaxation (diastole) is achieved when SERCA2a pumps Ca2+ back into the SR.

2+ The remaining Ca that entered the cell via Cav1.2 is exchanged across the plasma membrane for Na+ by the Na+/Ca2 + exchanger (NCX)23 (Figure 1B). This Ca2+ influx is responsible for the plateau phase of the cardiac action potential (Figure 1C).

2+ 24 Both RyR2 and RyR1 are activated by cytosolic Ca in the absence of Cav1.1 . This is pivotal for CICR in cardiac E-C coupling, but is also imperative for the appropriate execution of skeletal muscle E-C coupling because more than half of the RyR1 channels

2+ are not coupled to Cav1.1. It is generally believed that the skeletal muscle CICR Ca release amplifies the voltage-dependent Ca2+ release5,25. Interestingly, the amount of channels not coupled to Cav1.1 channels is not the same in all skeletal muscle types.

Slow-twitch skeletal muscles may have three or more uncoupled RyR1 channels for each

DHPR-linked RyR1 channel26-28. This may result from slow-twitch muscles having a greater CICR component, and this phenomenon may be related to the substantially slower rate of E-C coupling in slow-twitch muscle29.

Ryanodine receptors

The mechanism underlying intracellular Ca2+ release during E-C coupling was a topic of great debate in the 1970s and 1980s, with hypotheses ranging from the release of

2+ intracellular Ca by chemical transmission across the triadic junction through IP3 receptors30, by Na+/Ca2+ exchange31, or by a mechanical link between the t-tubule and an

SR Ca2+ release channel32. A breakthrough early in the 1980s was the identification, cloning and molecular characterization of the elusive SR Ca2+ release channel as a very high molecular weight protein with a high affinity for the plant alkaloid, ryanodine33-35.

Ryanodine is found naturally in the stem and roots of the plant, Ryania speciosa, which grows in Central and South America. This drug was being tested at that time as an insecticide due to its strong paralyzing effect on insects36. Subsequently, ryanodine was

used as a high affinity ligand to purify its receptor from SR preparations. Purified ryanodine receptors were shown to be Ca2+ release channels in skeletal37 and cardiac38 muscles, and ryanodine was shown to lock the channel open in a slow-gating subconductance state. Later studies determined that openings of the ryanodine-bound channel are long-lived and have a smaller unit current (∼1/3 or ∼1/2 of control amplitude) relative to open RyRs without the plant alkaloid bound. This observed Ca2+ release is the mechanism underlying the paralytic action of ryanodine5,34.

Ryanodine receptors are a family of Ca2+ release channels with large conductance for both monovalent and divalent cations (∼700 pS when K+ is charge carrier and ∼100 pS when Ca2+ is charge carrier) and relatively low Ca2+ selectivity (selectivity Ca2+/K+ ∼6)5.

Three isoforms of the mammalian ryanodine receptor have been identified and cloned,

RyR1, RyR2 and RyR3. The predominant isoform in skeletal muscle is RyR1 and in cardiac muscle it is RyR2. RyR1 and RyR2 play crucial roles for E-C coupling in these tissues5,39. RyR3 is also expressed in skeletal muscle at low levels40, though its physiological role in this tissue has not been identified. All three isoforms are expressed in the mammalian brain. RyR1 is enriched in Purkinje cells of the cerebellum, RyR2 is predominantly expressed in the dentate gyrus of the hippocampus41, and RyR3 has been detected in the hippocampal CA1 pyramidal cell layer, the basal ganglia, and olfactory bulbs42. RyR2 is also found in pancreatic islets43 and RyR1 and RyR3 have been reported in leukocytes44. All three isoforms are present in smooth muscle cells45, and a distinct gene encodes for each. RyR-encoding genes share an approximately 70% amino acid sequence identity46.

Numerous molecules, including Ca2+, Mg2+, and ATP, regulate the ryanodine receptor channel. Low intracellular [Ca2+] (1–10 µM) activates RyR and high intracellular [Ca2+]

(1–10 mM) inhibits channel activity. Though the effects of luminal Ca2+ on channel regulation have not been clearly defined, single channel studies show that luminal Ca2+

47 2+ affects single channel Po . The effects of Mg /ATP are more complex; cytosolic ATP is a RyR activator and cytosolic Mg2+ is a potent inhibitor48. However in the physiological state most ATP is in its Mg2+-bound form. Free ATP (∼300 µM in the cytosol) is the species that binds to and activates the RyR channel49. The effects of Mg2+ and ATP on ryanodine receptor channels are also isoform-specific; in the presence of physiological levels of Mg2+ and ATP, RyR1 requires less Ca2+ to activate than RyR2 or RyR350.

In addition, numerous pharmacological agents regulate RyR activity, and are thus useful as diagnostic tools to probe channel activity. Caffeine and 4-chloro-m-cresol (4-CmC) activate RyRs. These drugs are thought to facilitate channel opening by increasing the sensitivity of RyRs to Ca2+-dependent activation51. Ruthenium red is a commonly used

RyR blocker52. Local anesthetics such as procaine and tetracaine also block RyR channel activity in bilayers and in intact cells53,54.

The intracellular location of RyRs makes the study of their biophysical properties that much more complex; it is harder to patch clamp intracellular membrane-bound channels.

Three approaches have thus been undertaken to determine RyR activity: reconstitution of

the channel in planar lipid bilayers4, [3H]ryanodine binding55, and the measurement of

Ca2+ sparks56, a term used to describe elementary Ca2+ release events in the intact cell.

In order to determine the biophysical properties of single ryanodine receptor channels,

SR microsomes are purified and placed into a planar lipid bilayer chamber with two wells separated by a dividing wall with a small aperture painted with an artificial planar lipid bilayer. The voltage is clamped and the flow of current between the two chambers is measured in order to determine channel activity. Biophysical properties of the channel, such as conductance and ion selectivity as well as channel kinetics such as open probability, mean open and closed times and the channel reaction to small molecules and modulators can then be determined. The RyR-specific drugs ryanodine and/or ruthenium red are often applied at the end of each experiment to ensure that the measured current was the result of RyR channel activity5,24.

RyRs are relatively poorly selective for Ca2+ in vivo, and thus Ca2+ competes with Mg2+,

Na+, and K+ (intracellular concentrations of ∼1, 10, and 140 mM, respectively) for occupancy of the RyR channel pore. Thus, it should be noted that measurements of current from ryanodine receptor channels that have been reconstituted in planar lipid bilayers may reflect the lack of these in vivo competitors for the Ca2+ channel binding site57. Furthermore, it is standard when measuring single channel recordings to set the trans [Ca2+] much higher than the SR [Ca2+] in its native environment (53 mM trans vs. 1 mM in SR) to increase the driving force for Ca2+ when the channel opens. Thus measurements using single channel recordings are useful when making relative, not

absolute quantitative comparisons. In addition, the high trans [Ca2+] inhibits the opening of RyRs inserted in the opposite orientation in the bilayer lipid membrane due to inhibition of the channel by high cis [Ca2+].

When ryanodine receptor channels are in their open conformation, a high affinity ryanodine-binding site becomes exposed. The [3H]ryanodine binding technique makes use of this inherent property of ryanodine receptor channels. In these experiments ryanodine is pre-labeled with radioactive tritium [3H]. Purified RyR1 or isolated SR microsomes are, incubated with the [3H]ryanodine, and the excess [3H]ryanodine is washed. Preferential binding to the open state of RyRs provides an indirect measure of

RyRs activity55. Channel activators, such as µM [Ca2+] and ATP and caffeine increase the levels of bound [3H]ryanodine5,58.

In order to more effectively grasp the in vivo behavioral characteristics of ryanodine receptor channels, an alternate method to determine the biophysical properties has been developed thanks to advances in laser scanning confocal microscopy. RyR clusters normally remain closed until sarcolemmal depolarization and the activation of Ca2+ release and muscle contraction; however an individual cluster can occasionally open resulting in an elementary Ca2+ release event termed "Ca2+ spark"56,59. Ca2+ sparks are short lived local release events that can either be spontaneous or triggered by Cav1.1/

60 2+ 61 Cav1.2 activation . A Ca spark is generated by 4-6 RyRs and typically lasts approximately 50 ms and spreads to the size of about 2 µm in diameter, arising from a point source of Ca2+ in a ryanodine-sensitive manner59. The simultaneous generation of a

large number of Ca2+ spark release events causes the formation of the Ca2+ transient during physiological muscle excitation. However, increased Ca2+ spark activity has been observed in numerous pathophysiological states, such as osmotic shock and in dystrophic muscle62. It is thus possible to measure these Ca2+ release events using confocal microscopy, and increased Ca2+ spark release events are an indication of leaky ryanodine receptor channels.

Efflux of Ca2+ from the sarcoplasmic reticulum through ryanodine receptor channels under resting conditions, otherwise known as Ca2+ leak, affects the SR Ca2+ content and subsequently E-C coupling gain63. SR Ca2+ leak may occur as a result of various RyR1 modifications, as described below. Over time, this leak can lead to reduction in SR Ca2+ content, with less Ca2+ available for release and consequently weaker muscle contraction.

Furthermore, elevated resting cytosolic [Ca2+] may lead to several Ca2+-induced pathophysiological pathways, such as, transcriptional changes, activation of apoptotic pathways64 (Figure 2A-D) and membrane depolarization caused by electrogenic ion transport via NCX (leading to cardiac arrhythmia).

Ryanodine receptor regulation by post-translational modifications and regulatory proteins

Ryanodine receptor channels in the intracellular environment are ~2300-kDa homotetrameric complexes of four ~565-kDa subunits that form a central pore. Cryo- electron microscopy 3D reconstructions show that the RyR has an overall mushroom shape, with the mushroom stalk spanning the membrane and the cap residing in the

cytosol65. The large cytoplasmic N-terminal domain of RyRs modulates the gating of the channel pore, which is located on the C-terminus. The N-terminus forms a large cytosolic scaffold that interacts with accessory proteins, creating a regulatory macromolecular signaling complex46. When RyRs are solubilized, each component of the macromolecular complex co-sediments with the channel on sucrose density gradients66. Both, the components of the macromolecular channel complex and post-translational modifications of the channel are pivotal to local regulation of RyR activity.

The RyR macromolecular complex consists of cyclic AMP dependent protein kinase A

(PKA)66, which is bound to RyR via the anchoring protein, mAKAP (AKAP6)66; protein phosphatase 1, which is bound to RyRs via its targeting protein, spinophilin67; protein phosphatase 2A, which selectively associates with RyRs through its adaptor protein,

PR13068,69; Ca2+-dependent calmodulin kinase II (CaMKII)70,71, and the channel- stabilizing subunit, calstabin1 and calstabin2 in skeletal and cardiac muscle, respectively

(calcium channel stabilizing binding protein, also known as FK506-binding protein 12,

FKBP12 and FK506-binding protein 12, FKBP12.6 in skeletal and cardiac muscle, respectively)72. Leucine and isoleucine zippers (LZs and LIZs) are α-helical heptad repeats of Ile or Leu/Ile motifs that are involved in protein-protein interactions. RyRs were shown to bind adaptor/targeting proteins for regulatory kinases (PKA) and phosphatases (PP1 and PP2A) via conserved LZ/ILZ motifs73. This feature of the ion channel macromolecular complex is shared by intracellular Ca2+ release channels and voltage-gated channels74.

The catalytic and regulatory subunits of PKA, PP1, and the phosphodiesterase PDE4D3 regulate PKA-mediated phosphorylation of RyR1 at Ser2843 (Ser2844 in rodents), as determined using phosphopeptide mapping or site-directed mutagenesis of recombinant

RyR164,75-77. The role of PKA phosphorylation of RyR1 remains controversial, however, because some groups have found little or no effect of this post-translational modification on channel function78. Our laboratory has demonstrated that PKA-mediated phosphorylation of RyR1 at Ser2844 increases RyR1 sensitivity to cytoplasmic Ca2+, reduces the binding affinity of calstabin1 for RyR1, and destabilizes the closed state of the channel75,79. RyR1-S2844A mutant channels could not be PKA phosphorylated and did not show the same PKA-dependent increase in open probability. A RyR1-S2844D mutation that mimics PKA phosphorylation of the channel caused increased open probability80. Chronic PKA hyperphosphorylation of RyR1 at Ser2844 (defined as PKA phosphorylation of 3 or 4 of the 4 PKA Ser2843 sites present in each RyR1 homotetramer) results in leaky channels (channels prone to opening at rest), which contribute to the skeletal muscle dysfunction that is associated with persistent hyperadrenergic stress such as occurs in individuals with heart failure75,81. Furthermore, during exercise phosphorylation and other post-translational modifications of the RyR1 complex induce leaky channels and dissociation of calstabin1 from the channel complex, which plays a mechanistic role in limiting exercise capacity. Pharmacologically preventing calstabin1 depletion from the channel caused increased skeletal muscle force and exercise capacity72.

Depletion of PDE4D3 from the RyR1 complex contributes to PKA hyperphosphorylation of the RyR1 channel during chronic hyperadrenergic stress75. In rats with heart failure, the level of RyR1 PKA phosphorylation positively correlates with early fatigue in isolated soleus muscle during repetitive tetanic contractions75. Furthermore, preventing the dissociation of calstabin1 from the RyR1 complex and fixing the RyR1 leak in heart failure using the channel-stabilizing small molecule, S107, can improve muscle fatigue in mice with heart failure80.

Calstabins1 and 2 are immunophlins with peptidyl-prolyl-cis-trans isomerase activity82.

They share 85% sequence identity and form amphiphilic β-sheet structures that facilitate protein-protein interactions with ryanodine receptor channels. Calstabins are widely expressed and serve a variety of cellular functions. Each RyR tetramer binds four calstabin proteins (one molecule per monomer). Calstabin1 is expressed at higher levels in striated muscles83. RyR2 channels exhibit a higher affinity for calstabin2 in vivo83.

The binding of calstabins to RyRs may require that a peptidyl-prolyl bond is constrained in the high-energy transition-state intermediate between cis and trans. This hypothesis is supported by the fact that mutating the V2461 amino acid to a smaller and less sterically hindered glycine increases mobility around the peptidyl-prolyl bond, allowing for for cis/trans isomerization and subsequent disassociation of calstabin1 or calstabin2 from

RyR184. Furthermore, molecular modeling studies have demonstrated that the proline in the calstabin-binding region of RyR induces a bend in the alpha helix, which imposes a

twisted amide transition state on the peptidyl-proline bond and enables calstabins to bind to this domain85.

Physical and functional interaction between RyR1 channels and their associated calstabins are essential for the coordinated gating behavior that is necessary for the sharp rise in intracellular [Ca2+] during muscle contraction, termed coupled gating84. Coupled gating is the synchronous opening and closing (gating) of channels, which enables arrays of ryanodine receptor channels to gate in unison. This phenomenon both enhances the efficiency and propagates the appropriate and timely termination of SR Ca2+ release.

Coupled gating facilitates termination of SR Ca2+ release by inducing the channels to close in unison. Thus dissociation of calstabin1 from RyR1 causes stochastic gating of the channels79.

In addition to facilitating coupled gating, calstabin1 association with RyR1 affects other biophysical properties of the channel. Dissociation of calstabin1 from RyR1 increases channel open probability and induces subconductance states as observed in ryanodine receptor channels reconstituted in planar lipid bilayers86. Subconductance states are channel open events that are characterized by reduced unitary current amplitude.

Association of calstabin1 with the intracellular Ca2+ release channel stabilizes the closed state of the channel86.

Other members of the macromolecular complex regulate ryanodine receptor channel activity. The ubiquitously expressed Ca2+-binding protein, Calmodulin (CaM), binds

RyR1 at one per protomer stoichiometry. Apo-CaM (Ca2+ free) is a partial agonist at nanomolar Ca2+ concentrations, whereas at higher Ca2+ concentrations (greater than 1

2+ 87 µM), Ca -bound CaM functions as an inhibitor . CaM also modulates Cav1.1 gating in the t-tubule membrane, and Ca2+-CaM may inhibit binding of the C-terminal tail of the

88 2+ Cav1.1 α1S-subunit to RyR1 . Interestingly, CaM and the small dimeric Ca binding protein, S100A1, bind to an overlapping domain on RyR1 and it is thought that they function together to regulate channel function89.

Calsequestrin (CSQ) is located within cardiac, skeletal and smooth muscle SR and has also been described in other locations such as the brain (cerebellum). It is a high-capacity, low-affinity Ca2+ binding protein that forms a luminal sensor complex with triadin and junctin. The complex associates with RyRs through luminal regions of triadin and junctin90,91. CSQ also has the ability to directly modulate RyR function through Ca2+- dependent conformational changes, though the exact nature of the CSQ-RyR modulation has not been determined92.

RyR Redox sensing

Ryanodine receptors are tightly regulated by their redox state. Redox signaling relies on oxidants and antioxidants that react preferentially (nitrosylation and oxidation) with redox sensitive sulfhydryl groups of cysteine residues. Redox regulation of sulfhydryl groups has the potential to modify both the structure and function of proteins.

Each RyR1 tetramer contains 100 cysteines per monomer and of them 25–50 are accessible. Approximately twenty cysteine residues have been classified as available for redox modifications93, with six to eight of these categorized as hyperreactive, making them suitable specifically for modification by oxidation94. Moreover, maladaptive cAMP- dependent PKA-mediated phosphorylation and redox-dependent modifications of the

RyR1 have been linked to impaired Ca2+ handling and contractile dysfunction in chronic muscle fatigue, heart failure and muscular dystrophy46,72,81,95. Other components of E-C

96 coupling machinery are also regulated by their redox state, including SERCA and Cavs .

− The endogenous free radicals nitric oxide (NO•) and superoxide (O2 • ) are highly reactive redox agents with short half-lives that have the ability to modulate protein redox state. They may also be converted to non-radical species of lower reactivity and with longer half-lives such as S-nitrosoglutathione (GSNO) or hydrogen peroxide (H2O2) through enzymatic or non-enzymatic chemical reactions. Much of the cellular H2O2 is formed via the dismutation of superoxide by superoxide dismutase (SOD) enzymes97.

Reactive oxygen species (ROS) generation occurs at multiple sites in striated muscle including NADPH oxidase (NOX) associated with the plasma membrane or the SR/ER98,

99 coenzyme Q10 , xanthine oxidase (XO), lipoxygenase (LOX), phospholipase

100 101 A2 (PLA2) and complexes I and III of the mitochondrial electron transport chain

102 (Figure 3A). The relatively long cellular half-lives of H2O2 (~1 ms ) and GSNO relative

− 103 to their more reactive counterparts (the cellular half-life of O2 • is ~1-4 µs ) allow longer diffusion within cells and across cellular membranes, making them able to react

with many different cellular molecules, and activate a wide array of signaling pathways.

Due to their high levels of molecular stability, these less reactive oxidative agents play critical roles in signaling-related modification of cysteine residues.

Alterations in the redox state of ryanodine receptor channels can result in either activation104 or inactivation of the channel105. Both nitrosylation and oxidation affect skeletal muscle RyR1 function and subsequent Ca2+ signaling106,107. Ablation of the antioxidant enzyme superoxide dismutase 1 (SOD1) increases superoxide levels in murine skeletal muscle resulting in reduced specific force and accelerated age-dependent muscle pathology108. Increased levels of ROS in the aged muscle are associated with altered cellular Ca2+ handling108. Mice with a malignant hyperthermia mutation (Y522S) in RyR1 exhibit SR Ca2+ leak, which promotes mitochondrial dysfunction and oxidative stress-mediated changes of the RyR1109. Moreover, oxidation-dependent modifications of

RyR1 can increase SR Ca2+ leak.

A vicious cycle between RyR1-mediated leak of Ca2+ from the sarcoplasmic reticulum and mitochondrial ROS production has been proposed based on these data109. This vicious cycle hypothesis is particularly compelling due to the presence of protein tethers connecting mitochondria and SR/ER110, which mediate functional association as well as redox coupling between skeletal muscle mitochondria and SR111. SR/ER fragments remain associated with the mitochondria after fractionation, which allows the isolated

SR/ER–mitochondrial complexes to sustain local Ca2+ communication112. This may

provide an anatomical basis for the proposed vicious cycle between redox regulation of/by the mitochondria and SR.

Oxidation of critical sulfhydryls located on the cytoplasmic side of ryanodine receptor channels affect both the gating properties of the channels and responsiveness to channel modulators such as adenine nucleotides, caffeine, Ca2+ and Mg2+ sensitivity, the channel's ability to bind calmodulin113 and the association of calstabin1 with RyR1114. Likewise, cysteine nitrosylation of RyR1 increases channel activity115,116 and dissociates calstabin1 from the intracellular Ca2+ release channel, resulting in leaky channels, reduced SR

Ca2+ release upon stimulation and a subsequent decrease in muscle function95.

Ryanodine receptor channels are also highly susceptible to modification by other endogenous redox agents, including glutathione (GSH), glutathione disulfide (GSSG),

NADH, and by changes in the GSH/GSSG ratio117. S-glutathionylation of RyR1 reduces its affinity to calmodulin114. In addition to the direct effects of these endogenous redox agents, there are also indirect effects of oxidative agents on RyR1 function. For example,

NO-dependent guanylyl cyclase–mediated generation of cGMP can activate protein kinase G, which, in turn, can phosphorylate RyR1118. This is especially relevant since NO have been found endogenously nitrosylate RyR1118. Moreover, reactive nitrogen species can substantially modulate catalase and other antioxidant enzymes in skeletal muscle119-

121, and the C3635 or other cysteine residues of RyR1 may undergo selective covalent modification by NO, thereby contributing to redox regulation of the ryanodine receptor macromolecular complex122. Thus, interplay between the endogenous oxidative agents is

pivotal to redox regulation of ryanodine receptors.

Though they may become maladaptive, redox modifications are necessary for appropriate

RyR1 function; RyR1 cysteines have been demonstrated to be S-nitrosylated, S- glutathionylated, and oxidized under distinct physiological conditions123,124. This apparent necessity for oxidative balance may help explain the elaborate and evolutionarily conserved interplay between the anti-oxidative and oxidative systems that regulate cellular redox state125,126. Namely, the mammalian antioxidant network consists of enzymes, such as catalase, glutathione peroxidase (GPx), thioredoxin reductases

(TRxs), superoxide dismutases (SODs), and soluble antioxidants such as glutathione

(GSH) and vitamin E.

The integrity of the oxidative/anti-oxidative systems is particularly significant in aging.

Increased free radical generation and the resulting oxidative damage accumulated in organisms are likely to be involved, in the progression of abundant age-related diseases.

Recently there have been efforts to study mitochondria-derived free radicals in health- and lifespan by experimentally expressing catalase in the mitochondria using in vitro models127, adeno-associate viral vectors (AAV)128 and most recently by genetically engineering its overexpression in mice129. These transgenic mice, MCat mice, in which the human catalase is overexpressed and retargeted from the peroxisome to the mitochondrial matrix, display 10-20% increase in maximum and median lifespan 129, exhibit age-related insulin resistance130, attenuated energy imbalance131 and demonstrate myocardial protection against heart failure132,133.

RyR1-related skeletal muscle pathophysiology

Ryanodine receptor channel mutations and post-translational modifications have the propensity to cause numerous skeletal muscle pathologies. Malignant hyperthermia (MH) is a pharmacogenetic shock syndrome that may be triggered by halogenated inhalation anesthetics (e.g., halothane) and/or depolarizing muscle relaxants (e.g., succinylcholine)46. Genetically susceptible patients exposed to these drugs develop skeletal muscle rigidity, rapidly rising body temperature, fast heart rates, and acidosis. In

MH-susceptible patients, three mutation clusters have been identified in the RyR1 gene: the N-terminal region C35-R614, the central region D2129-R2458, and the C-terminal region I3916-A4942. RyR1 MH mutations result in abnormal sensitivity to channel activation by caffeine, halothane, 4-chloro-m-cresol and membrane depolarization134. The muscle relaxant drug, dantrolene, is effective in preventing MH crises due to its ability to inhibit SR Ca2+ leak via RyR1135,136. Although RyR1 mutations account for over 50% of all MH cases, other MH susceptibility loci exist, including Cav1.1 missense mutations of

137 a conserved arginine residue in the Cav1.1 III-IV linker region .

Another characteristic RyR1 dysfunction-related disease is central core disease (CCD).

Unlike MH, which presents in otherwise healthy individuals, CCD manifests as proximal muscle weakness starting at infancy with delayed attainment of motor milestones and lifetime musculoskeletal dysfunction. CCD has been linked to mutations in the RyR1 gene in the same regions as MH. Furthermore, CCD patients are frequently susceptible to

MH138,139. Although variable, skeletal muscle biopsies reveal amorphous cores or regions

devoid of mitochondria and oxidative enzyme activity46. RyR1 mutations linked to CCD alter RyR1-dependent Ca2+ release mechanisms140, cause SR Ca2+ leak141, and have the propensity to cause E-C uncoupling142.

Muscular Dystrophy (MD) is a progressive multisystem genetic disease characterized by peripheral weakness and degeneration of both skeletal and cardiac muscles, leading to impaired locomotion, cardiomyopathy and early death. The dystrophin-glycoprotein compex (DGC) is a large complex of membrane-associated proteins that maintains skeletal muscle fiber integrity and protects muscle cells from mechanical, contraction- induced muscle damage143 (Figure 4). Among the DGC components, mutations in dystrophin and the sarcoglycans have been linked to muscular dystrophies, including

Limb-Girdle Muscular Dystrophies (LGMD), which may be caused by loss of the sarcoglycans. Duchenne muscular dystrophy (DMD), the most common X-linked disorder, affects 1 in 3,500 male births and typically results in death as a result of respiratory or cardiac failure by age 30. It is caused by loss of dystrophin, which leads to disruption of the DGC in the sarcolemmal membrane that connects the cytoskeleton and contractile apparatus of muscle cells to the extracellular matrix and plasma membrane144.

A contributing factor to the musculoskeletal degeneration in DMD is a structural and functional defect in RyR1 that leads to altered Ca2+ homeostasis in dystrophic muscle.

RyR1 channels in mdx mice are leaky, hypernitrosylated, and depleted of calstabin1.

Preventing calstabin1 depletion from RyR1 with S107, a compound that binds the ryanodine receptor channel and enhances the binding affinity of calstabin1 to the channel,

inhibits sarcoplasmic reticulum Ca2+ leak, reduces biochemical and histological evidence of muscle damage, improves muscle function and increases exercise performance in mdx mice, strongly suggesting a causative role for ryanodine receptor channel dysfunction in the mdx phenotype95.

A hallmark of aging is sarcopenia; the progressive decline in skeletal muscle function, characterized by reduced force generating capacity and loss of muscle mass 145,146.

Moreover, age-dependent deterioration of muscle function is not restricted to mammals as it is also observed in many animals including the nematode Caenorhabditis elegans

(C. elegans) 147. Indeed, loss of muscular strength is highly predictive of frailty, disability and mortality with increased age 148-150. Progressive development of muscle dysfunction is common in aged humans and in animal models of aging 150, and affects as many as 30-

50% of 80 year olds leading to profound loss of function in the elderly 146. Much attention has been focused on understanding how to reverse age-related muscle wasting, but there are no established treatments for sarcopenia at this time 150-152. In contrast, improving specific force production, which is also significantly reduced in aged muscle

153,154, has received significantly less attention.

Excitation-contraction coupling in C. elegans

C. elegans is a saprophytic nematode species that inhabits soil and leaf-litter environments in many parts of the world155. Sydney Brenner's seminal paper in 1974156 on the genetics of C. elegans introduced it as an important experimental model. The use of the nematode as a model to study physiological processes has an array of advantages,

including a fully sequenced genome, short lifespan and the ability to perform genetics on the organism. Thus, C. elegans has been successfully utilized in aging studies157 and as a tool for drug screening158,159. Numerous important discoveries in neuroscience, development, signal transduction, cell death, aging, and RNA interference (RNAi) have been made in the relatively short time since Brenner’s publication160.

Since the first gene that modulates lifespan was discovered in C. elegans in 1988161 many mutant nematode strains with alterations in lifespan have been developed, and it has become an important model used to study of the genetics of aging162,157. Though less extensively studied, aged nematode physiology conveys similarities to mammals that present it as a highly tractable model system in which to conduct aging studies.

Specifically, C. elegans exhibit age-dependent muscle deterioration and reduced locomotion. There is a reduction in muscle cell size associated with loss of cytoplasm and myofibrils, and progressive myofibril disorganization147. The nematode aging phenotype is strikingly similar to the mammalian aging phenotype, which includes muscle wasting and reduced force generating capacity (akin to the progressive defects in locomotion in the nematode)145,147,163.

E-C coupling in C. elegans has not been fully characterized, though there are key similarities between the vertebrate and nematode systems, both in terms of molecular structure and function. The body wall muscle of C. elegans consists of 95 muscle cells, which are arranged as staggered pairs in four longitudinal bundles located in four quadrants. Three of these bundles contain 24 cells each, whereas one bundle contains 23

cells. The body wall muscle of C. elegans, as in all other nematodes, is obliquely striated.

The vertebrate Z-disc (Zwischenscheibe disc) analog in C. elegans muscle is the dense body (DB), which functions to anchor and align thin filaments in striated muscle. Thick filaments are attached to M-line analogs (the middle of the H bands in vertebrates, where each myosin rod is joined end-to-end with its myosin rod neighbor). Both the DBs and the M-line analogs extend the entire depth of the lattice and anchor all filaments to the cell membrane and the underlying hypodermis and cuticle. Unlike other organisms where neurons send processes to their target muscle cells to make synapses, neuromuscular junctions of C. elegans are made by arms grown from muscle cells toward motor neurons164,165.

Although the nematode body wall muscle filaments are oriented parallel to the longitudinal axis of the muscle cell, adjacent structural units (M-lines and DBs) are offset from one another by more than a micron, rather than being in register as in vertebrate cross-striated muscle166. Therefore, the observed A–I striations occur at an angle of 5–7° with respect to the longitudinal axes of the filaments and the muscle cell, in comparison to 90° in vertebrate muscle. This arrangement of the sarcomeres likely creates a more evenly distributed muscle force application over the basal lamina and cuticle, allowing for smooth bending of the body rather than kinking167.

C. elegans SR consists of a network of vesicular membranous organelles surrounding the myofilament lattice. The flattened vesicles of SR extend around dense bodies and lay adjacent to the hypodermal side of the plasma membrane underneath the lattice, where

they are localized randomly between DBs 168. A gap of 12-14 nm separates the SR vesicles from the plasma membrane. No equivalent to the t-tubule system exists in C. elegans, possibly due to the direct apposition of SR to the plasma membrane168.

Following excitatory (cholinergic) neurotransmission at the neuromuscular junctions of C. elegans, opening of nicotinic acetylcholine receptor (ligand-gated ion channels) on muscle membrane is thought to initiate graded action potentials in muscle arms, which then converge and propagate to the contractile compartment of the muscle169-171.

C. elegans action potentials are all-or-none events with a stereotypical waveform. Action potential-induced elevations in intracellular [Ca2+] require the function of both, EGL-19

172 (homologous to the alpha 1 subunit of the mammalian Cavs ) and UNC-68 (homologous to mammalian RyRs)169,173. There are no voltage-activated Na+ channels in C. elegans so any extracellular ionic influx during E-C coupling is likely mediated by Ca2+ via EGL-

19169. The homolog of the mammalian T-type Ca2+ channel alpha 1 subunit, CCA-1, has also been implicated in shaping and/or generating action potentials by inward currents.

Furthermore, the voltage-gated K+ channel, SHK-1, and the Ca2+/Cl--gated K+ channel,

SLO-2, carry sustained outward K+ currents173. Though it has been suggested that the nematode E-C coupling progresses through Ca2+-induced-Ca2+-release, the relative mechanistic roles of EGL-19 and UNC-68 in nematode E-C coupling have not been established173.

Null egl-19 mutants are not viable, and gain-of-function mutations cause prolonged muscle action potentials and delayed muscle relaxation (known as myotonia).

Furthermore, reduction-of-function mutations cause a reduced rate of depolarization and feeble contractions172. In the pharyngeal muscles, cca-1 null mutants are missing one of the two components of the inward Ca2+ current, the low voltage activated (LVA) current, suggesting that this channel mediates the LVA, and that EGL-19 channel carries the high voltage activated (HVA) current in the pharyngeal muscle174. cca-1 mutants exhibit pharyngeal pumping defects175.

unc-68 is the only RyR gene in the C. elegans genome (Figure 5A) 176. It is expressed in body wall muscles, the terminal bulb muscle of the pharynx, vulval and uterine muscles, diagonal muscles of male tail, and the anal sphincter and depressor muscles177. unc-68 null mutants exhibit locomotive defects characterized by the “unc”, or “uncoordinated” phenotype156. This null mutation also confers visually detectable defects in muscle function and motility. Additionally, this mutation induces defective pharyngeal pumping. unc-68 null mutants also exhibit slower growth rate and reduced number of offspring177.

Within the body wall muscle, UNC-68 is thought to be localized to SR vesicles, primarily between the rows of dense bodies in the A-band region177. Treatment with ryanodine, induces contractile paralysis in wild-type C. elegans, but null unc-68 mutants are unaffected by this pharmacological intervention156,176,178. Ca2+ transients triggered by action potentials in C. elegans body wall muscles require UNC-68173,178 and the nematode ryanodine receptor channel shares 40% sequence identity with human RyR1179.

Furthermore, important functional domains are conserved including a PKA phosphorylation site and two canonical Ca2+ binding EF-hand motifs180.

The C. elegans genome contains eight FK506-binding proteins with varying levels of sequence identity compared to mammalian calstabins (Figure 5B). The one with the highest homology to a mammalian isoform is FKB-2. FKB-2 exhibits ~60% sequence identity to mammalian calstabins181, however little is known about its function. In general there are few reports on the function of FKBs in C. elegans; FKB-3, 4 and 5 are reported to be involved in nematode development, collagen biogenesis, and the formation of an intact exoskeleton under adverse physiological conditions182,183. FKB-4 is necessary for dietary restriction-induced increased nematode lifespan184. Furthermore, FKB-6, which is the sole TRP-containing immunophilin in the C. elegans genome, interacts with C- terminal C. elegans DAF-21. This finding is consistent with the interaction found between the large human immunophilin TPR domains and human Hsp90185.

C. elegans body wall muscle also has a unique feature in that the rhythmic defecation cycle that involves posterior body wall muscle contraction is regulated by protons acting as a direct transmitter from intestinal cells to stimulate muscle contraction. During the C. elegans defecation motor program the posterior body muscles contract even in the absence of neuronal inputs. The release of caged protons is sufficient to induce muscle contraction in this area of the body wall muscle186. Though there are significant similarities between the nematode and mammalian systems, it is therefore important also to note the differences.

Redox balance/imbalance in aging C. elegans

The molecular machinery underlying the oxidative/anti-oxidative systems in C. elegans has not been fully characterized, though mitochondria are considered the major source of cellular ROS187 (Figure 3B). In addition, the C. elegans genome encodes two NOX homologs, BLI-3 and DUOX-2188, however they have not been implicated in ROS signaling; BLI-3 functions in tyrosine cross-linking in the extracellular matrix189.

Furthermore, their mammalian NOX homologs, which are dual oxidases, play a role in the biosynthesis of thyroid hormones in the extracellular matrix190. It is possible that further investigation will reveal a more elaborate oxidative system in the nematode.

Numerous components of the anti-oxidative system in C. elegans have been identified

(Figure 3B). The mitochondrial thioredoxin reductase, TRXR-2, is expressed in nematode muscle cells (body wall and pharyngeal muscles) and TRXR-2 deficiency causes defective longevity and delayed development under stress conditions191. 2-Cys peroxiredoxin 2 (PRDX-2) is a highly abundant protein that constitutes ~0.5% of the total C. elegans proteome and is an important part of the nematode H2O2 detoxification system192. Deletion of prdx-2 increases the sensitivity of C. elegans toward exogenous peroxide treatment and causes a significant decrease in the lifespan of these worms, especially at lower cultivation temperatures. Several genes in the C. elegans genome encode SOD enzymes: sod-1 encodes cytosolic CuZn-SOD193, sod-2 and sod-3 each encode Mn-SOD194-196, and sod-4 encodes extracellular CuZn-SOD197. The functional differences between the two Mn-SODs in C. elegans are not yet known. Furthermore, two catalase genes are present in C. elegans. ctl-1 encodes a cytosolic catalase; ctl-

2 encodes a peroxisomal catalase. A ctl-1 mutation was shown to suppress the life extension phenotype of daf-2, age-1, and clk-1 mutants198.

Rationale and aims

The overarching goal of this study was to examine RyR1 regulation in health and disease.

We hypothesized that regulation of ryanodine receptors by post-translational modifications plays a mechanistic role common to both the physiological and pathophysiological states in skeletal muscle. More specifically, we hypothesized that similar post-translational modifications of RyR1 may contribute to the enhancement and determent of contractile function. We addressed this hypothesis by investigating regulation of RyR1 in skeletal muscle inotropy and the role and regulation of RyR1 in

Limb-girdle muscular dystrophy (LGMD) in two distinct murine mouse models in aim 1.

We chose to study LGMD because a null mutation in one of the sarcoglycans results in loss of the whole sarcoglycan complex but not of dystrophin199. However mutations in dystrophin, which cause DMD, also lead to loss of the sarcoglycans200. This led us to hypothesize that the loss of sarcoglycans is the central upstream event in DGC-associated muscular dystrophies.

Secondly, we hypothesized that common mechanisms regulate skeletal muscle weakness in Muscular Dystrophy and aging, and we thus addressed the role and source of RyR1 oxidation in aged mice. Finally, in aim 3, we hypothesized that the classic aging model,

C. elegans, would be an excellent model in which to study of the molecular determinants of muscle physiology in aging.

Aim 1: To determine if post-translational modifications such as oxidation and phosphorylation of RyR1 underlie skeletal muscle inotropy and muscular dystrophy. We hypothesize that regulation of ryanodine receptors by post-translational modifications plays a mechanistic role in both the physiological and pathophysiological states in skeletal muscle. To test this hypothesis, we investigated the role of post- translational modifications of RyR1 in skeletal muscle inotropy and Limb-girdle muscular dystrophy.

Aim 2: To determine the mechanistic role and source of RyR1 oxidation in age- dependent skeletal muscle weakness. A common phenotype of muscular dystrophies and aging is a progressive decline in skeletal muscle function. Defective SR Ca2+ release has been reported in age-dependent muscle weakness201,202 and maladaptive modifications of the RyR1 macromolecular complex have been implicated 203. However, the mechanism underlying impaired SR Ca2+ release in aging muscle had not yet been elucidated. We therefore hypothesized that an underlying mechanism of age-dependent skeletal muscle weakness is RyR1 dysfunction.

Aim 3: To elucidate the regulation of the C. elegans RyR homolog, UNC-68, in physiology throughout nematode lifespan. Strikingly, despite an approximately 2,000- fold difference in the lifespans of humans and C. elegans204, both exhibit age-dependent reductions in muscle function and motor activity that ultimately contribute to senescence and death. Due to its short lifespan and well-characterized genome, C. elegans has been

used as a model for the study of the genetics of aging and lifespan determination157,162 including the age-dependent decline in locomotion147,205,206. Age-dependent reduction in locomotion in C. elegans has been attributed to degeneration of the nervous system207 and the body wall musculature147,208. We hypothesized that UNC-68 is regulated in young and aged worms by FKB-2, the FKBP in the C. elegans genome with the highest sequence identity to mammalian calstabins.

Figure 1: Cardiac vs. skeletal muscle E-C coupling. (A) Schematic representation of skeletal muscle E-C coupling; (B) Schematic representation of skeletal (solid line) vs. cardiac (dashed line) action potentials; (C) Schematic representation of cardiac muscle E- C coupling; Major differences between the two processes depicted are direct physical contact between Cav1.1 and RyR1 vs. CICR in cardiac muscle and NCX in cardiac muscle.

Figure 2: Physiological vs. pathophysiological skeletal muscle E-C coupling. (A) Schematic representation of physiological E-C coupling; (B) Schematic representation of pathophysiological E-C coupling, during which Calstabin1 (Cal1) has dissociated from RyR1, causing intracellular Ca2+ leak (dashed line), depletion of the SR Ca2+ store, and perturbed contraction (solid line); (C) Schematic representation of action potential (mV; solid line), muscle force (dashed line) and Ca2+ transient (grey line) in physiological and (D) pathophysiological contraction.

Figure 3: Oxidative systems in mammalian skeletal muscle and C. elegans. (A) Schematic representation of potential mechanisms for generation of reactive oxygen species in mammalian skeletal muscle. CAT, catalase; CuZnSOD, copper, zinc superoxide dismutase (SOD1); ecSOD, extracelluar superoxide dismutase (SOD3); GPx, glutathione peroxidase 1; eNOS, endothelial nitric oxide synthase (NOS3); NO, nitric oxide; MnSOD, manganese superoxide dismutase (SOD2); nNOS, neuronal nitric oxide synthase (NOS1); PM oxido-reductase, plasma membrane-located oxido-reductase; PLA2, phospholipase A2; VDAC, voltage-dependent anion channel; GPx, glutathione peroxidase; (B) Schematic representation of potential mechanisms for generation of reactive oxygen species in C. elegans.

Figure 4: Schematic representation of the dystrophin glycoprotein complex (DGC). The DGC spans the sarcolemmal membrane, linking the actin-based cytoskeleton to the overlying basal lamina. Transmembrane components of the DGC include sarcoglycans, sarcospan (SPN), and beta-dystroglycan. The cytoplasmic components of the complex include dystrophin, alpha-dystrobrevin, and syntrophins, the latter of which anchors the signaling molecule neuronal nitric oxide synthase (nNOS).

Figure 5: Sequence identity between mammalian and C. elegans ryanodine receptors and immunophillins. (A) Sequence identity (ID) and % positives (Pos) between mammalian RyRs and C. elegans homolog, UNC-68; (B) Sequence identity (ID) and % positives (Pos) between mammalian calstabins and C. elegans homologs (FKB1- 8).

Chapter 2: Post-translational modification of the ryanodine receptor type 1 underlies skeletal muscle weakness in the physiological and pathophysiological state

Chapter 2, Section 1: Phosphorylation of the ryanodine receptor by protein kinase

A is a molecular mechanism underlying fast twitch skeletal muscle inotropy.

Skeletal muscle displays increased force upon adrenergic stimulation 209,210, however, despite the well-established role of this physiological response, the molecular mechanism is not known. Under stress conditions, stimulation of Gs-protein coupled β-adrenergic receptors (β1 and β2-receptors) activates adenylyl cyclase and causes generation of the second messenger cAMP, which in turn activates cAMP-dependent protein kinase A

(PKA; Figure 6)211,212. PKA-mediated phosphorylation of Ca2+ handling proteins is a key mechanism underlying increased contraction of cardiac myocytes 213,214.

In skeletal muscle, unlike in cardiac muscle, contraction does not depend on Ca2+ entry across the plasma membrane and the Ca2+ needed to activate the myofilaments comes from the release of SR Ca2+ 215. Fast-twitch skeletal muscle fibers (type II) lack the sarcoplasmic reticulum (SR) Ca2+ ATPase (SERCA2a)-associated protein phospholamban, a target of PKA phosphorylation that regulates SR Ca2+ load in cardiomyocytes. β-receptor stimulation in the absence of phospholamban does not lead to changes in SERCA activity or SR Ca2+ load in fast-twitch muscles 216,217. Furthermore, neither myofilament Ca2+ sensitivity nor sarcolemmal Ca2+ fluxes are affected by β- adrenergic activation in fast twitch skeletal muscle 216. Nonetheless, fast twitch muscle

fibers display a robust inotropic response to β-receptor agonists due to potentiated increase in cytoplasmic [Ca2+]218.

Using phosphopeptide mapping or site-directed mutagenesis of recombinant RyR1, our laboratory75 and others219, have identified the PKA phosphorylation site of human RyR1 as RyR1-S2843 (RyR1-S2844 in rodents). Here, we used a novel genetically altered mouse (RyR1-S2844A) that expresses a mutant RyR1 that cannot be PKA phosphorylated to show that β-receptor agonist-dependent increase in twitch [Ca2+] and force in mammalian fast-twitch skeletal muscle are dependent on phosphorylation of a single amino acid, serine 2844 in RyR1.

Chapter 2, Section 1 Results

To examine the role of PKA-dependent RyR1 phosphorylation in skeletal muscle inotropy we generated a mutant mouse (RyR1-S2844A) in which the serine at the RyR1

PKA phosphorylation site was substituted with alanine, which cannot be phosphorylated

75. There was no difference in body weight between the WT and RyR1-S2844A mice

[WT, 28±1.1 g, n=8; RyR1-S2844A, 26±1.3 g, n=9; mean ± SEM, P = n.s. (unpaired t- test)]. Force generation in isolated fast twitch extensor digitorum longus (EDL) muscles from the RyR1-S2844A mice was compared to wild-type (WT) mice. At baseline, force- generating capacity in the RyR1-S2844A and WT EDL was not different, as evidenced by the similar force-frequency curves (Figure 7).

Brief application of the β-receptor agonist, isoproterenol (ISO; 1 µM, 15 min), to WT

EDL muscles caused a significant increase in twitch force (Figure 8A-B). In contrast,

EDL muscles isolated from the RyR1-S2844A mouse did not display an increase in twitch force in response to ISO treatment (Figure 8C-D). These data indicate that RyR1-

S2844 phosphorylation plays a key role in the increase in peak force in fast twitch skeletal muscle following adrenergic stimulation.

Muscle force is graded by SR Ca2+ release, which determines cytoplasmic [Ca2+], and β- adrenergic agonists have been shown to increase cytoplasmic [Ca2+] in fast-twitch flexor digitorum longus (FDB) muscle216. However the mechanism underlying this inotropic response remains obscure. To examine ISO induced enhancement of twitch Ca2+, isolated

FDB muscle fibers were loaded with the cell permeable fluorescent indicator for Ca2+, fluo-4 and twitch contractions were triggered by electrically stimulating the fibers220.

Changes in fluo-4 fluorescence were measured, in the same muscle fibers under control conditions and after 10 min in the presence of ISO (1µM). In WT, the peak fluo-4 signal increased significantly following ISO treatment (Figure 9A and C). Conversely, FDB muscle from RyR1-S2844A knock-in mice did not demonstrate a change in fluo-4 peak fluorescence following ISO stimulation (Figure 9B and C).

To confirm the phosphorylation state of the Ca2+ release channel after ISO treatment, immunoprecipitated RyR1 from EDL muscle homogenates were immunoblotted using an

RyR1 PKA phosphorylation site (RyR1-S2844) epitope-specific antibody220. ISO treatment of WT muscles led to increased RyR1 phosphorylation, whereas no such

increase was detected in the RyR1-S2844A skeletal muscles (Figure 10A). In support of this finding using RyR1 isolated from RyR1-S2844A skeletal muscles we previously reported that heterologously expressed RyR1-S2844A channels cannot be PKA phosphosrylated75. We have previously reported that under conditions of chronic adrenergic drive, such as in heart failure, excessive PKA phosphorylation of the skeletal muscle RyR1 is associated with depletion of calstabin1 75,221. We therefore measured binding of calstabin1 to RyR1 in WT and S2844A muscle following acute ISO treatment.

Under these experimental conditions, calstabin1 binding was unaltered in both of the groups (Figure 10A).

Finally, to determine the effect of the RyR1-S2844A mutation on skeletal muscle strength in vivo, we measured forelimb grip strength in WT and RyR1-S2844A mice95.

The RyR1-S2844A muscle displayed reduced grip strength (Figure 10B).

Chapter 2, Section 1 Summary

Based on these data we have reported that PKA phosphorylation of a single amino acid of the RyR1, serine 2844, is a key signal in the physiological stress-induced inotropic response in skeletal muscle. Specifically, β-receptor agonist-dependent enhanced muscle

Ca2+, force and in vivo muscle strength responses following ISO stimulation were abrogated in RyR1-S2844A mice in which the serine in the PKA site in RyR1 was replaced with alanine. These data suggest that the molecular mechanism underlying skeletal muscle inotropy requires enhanced SR Ca2+ release due to PKA phosphorylation

of S2844 in RyR1. These findings elucidate a fundamental mechanism regulating physiological skeletal muscle function.

Chapter 2, Section 1 Methods

Muscle function

Extensor digitorum longus (EDL) muscles were dissected from hind limbs. Stainless steel hooks were tied to the tendons of the muscles using nylon sutures and the muscles were mounted between a force transducer (Harvard Apparatus) and an adjustable hook. The muscles were immersed in a stimulation chamber containing O2/CO2 (95/5%) bubbled

Tyrode solution (in mM: NaCl 121, KCl 5.0, CaCl2 1.8, MgCl2 0.5, NaH2PO4 0.4,

NaHCO3 24, EDTA 0.1, glucose 5.5). The muscle was stimulated to contract using an electrical field between two platinum electrodes (Aurora Scientific). At the start of each experiment the muscle length was adjusted to yield the maximum force. Isometric twitch force was produced by stimulating the muscle using a 0.5 ms pulse at supra-threshold voltage. Force was measured from each muscle before and after application of ISO (1

µM; 15 min). At the end of force measurement, the length (L0) and weight of the muscle was measured and the muscle was snap frozen in liquid N2.

Ca2+ imaging using fluorescent indicators in FDB muscle fibers

Single FDB fibers were obtained by enzymatic dissociation as previously described 222.

FDB muscles from both hind limbs were incubated for ~2 hours at 37ºC in ~4 ml

Dulbecco’s Modified Eagles Medium (DMEM) containing 0.3% collagenase 1 (Sigma) and 10% fetal bovine serum. The muscles were transferred to a culture dish containing

fresh DMEM (~4 ml) and gently triturated using a 1000 µl pipette until the muscles were dissociated. The cell suspension was stored in an incubator at 37ºC/5% CO2 until the start of the experiment.

FDB fibers were loaded with the fluorescent Ca2+ indicator Fluo-4 AM (5µM,

Invitrogen/Molecular probes) for 15 min at RT. The cells were allowed to attach to a laminin coated glass cover slip that formed the bottom of a perfusion chamber. The cells were then superfused with tyrode solution (in mM: NaCl 121, KCl 5.0, CaCl2 1.8, MgCl2

0.5, NaH2PO4 0.4, NaHCO3 24, EDTA 0.1, glucose 5.5; bubbled with O2/CO2 (95/5%)).

The fibers were triggered to twitch contraction using electrical field stimulation (pulses of

0.5 ms at supra-threshold voltage) and Fluo-4 fluorescence was monitored using a confocal microscope system (Zeiss LSM 5 Live, 40x oil immersion lens, excitation wavelength was 488 nm and the emitted fluorescence was recorded between 495-525 nm). The use of the single excitation/emission dye fluo-4 necessitates normalizing to pre- stimulation values to negate possible differences in dye loading and excitation strength.

Only fibers that were attached to the bottom of the perfusion chamber throughout the twitch stimulation were measured from. All experiments were performed in RT (~20ºC).

RyR1 immunoprecipitation and Immunoblotting

EDLs were isotonically lysed in 0.5 ml of a buffer containing 50 mM Tris-HCl (pH 7.4),

150 mM NaCl, 20 mM NaF, 1.0 mM Na3VO4, and protease inhibitors. This muscle lysate was used for either immunoblotting with CREB/pCREB antibodies or immunoprecipitation of the RyR1. For the immunoprecipitation, an anti-RyR antibody (4

µg 5029 Ab) was used to immunoprecipitate RyR1 from 250 µg of tissue homogenate.

The samples were incubated with the antibody in 0.5 ml of a modified RIPA buffer (50 mM Tris-HCl pH 7.4, 0.9% NaCl, 5.0 mM NaF, 1.0 mM Na3VO4, 1% Triton- X100, and protease inhibitors) for 1 hour at 4°C. The immune complexes were incubated with protein A Sepharose beads (Sigma, St. Louis, MS) at 4°C for 1 hour and the beads were washed three times with buffer. Proteins were separated on SDS-PAGE gels (6% for

RyR1, 15% for calstabin1) and transferred onto nitrocellulose membranes for 1 hour at

200 mA (SemiDry transfer blot, Bio-Rad). After incubation with blocking solution

(LICOR Biosciences, Lincoln NE) to prevent non-specific antibody binding, immunoblots were developed with anti-RyR (Affinity Bioreagents, Bolder, CO 1:2,000), anti-phospho-RyR1-pSer2844 (1:5000). All immunoblots were developed and quantified using the Odyssey Infrared Imaging System (LICOR Biosystems, Lincoln, NE) and infrared-labeled secondary antibodies.

Grip strength

Forelimb grip strength was measured in stressed WT and S2844A mice using an automated grip strength meter (BIOSEB in vivo research instruments, Vitrolles, France).

Mouse forelimbs were allowed to grab onto a metal grid (attached to a force transducer).

Thereafter, mice were pulled by the tail until losing grip. Maximum force was recorded.

The grip strength was calculated from the average of 3 repeated measurements in each mouse. This procedure activated stress signaling, as the mice resisted and displayed spontaneous defecation and/or urination. All measurements were performed blinded with respect to the mouse genotype.

Chapter 2, Section 2: Post-translational modifications of the ryanodine receptor mechanistically underlie skeletal muscle weakness in Limb-girdle muscular dystrophy

Muscular dystrophies (MD) comprise a group of inherited disorders affecting striated muscles that are characterized by progressive weakness and muscle degeneration. The dystrophin-glycoprotein complex (DGC) is a macromolecular structure of membrane- associated proteins that includes dystrophin and the sarcoglycan proteins (α-, β-, δ-, and

γ-sarcoglycan), which maintain fiber integrity and protect from contraction-induced muscle damage143,223 (Figure 4). Mutation-induced disruption of sarcoglycan proteins leads to limb-girdle muscular dystrophy (LGMD)224. A null mutation in one of the sarcoglycans results in loss of the whole sarcoglycan complex but not of dystrophin199.

However mutations in dystrophin, which cause the most common form of muscular dystrophy, Duchenne muscular dystrophy (DMD), also lead to loss of the sarcoglycans200. This implicates the loss of sarcoglycans as the central upstream event in muscular dystrophies.

Our laboratory has previously determined that an underlying mechanism of the musculoskeletal degeneration in DMD is RyR1 dysfunction. RyR1 channels in mdx mice are leaky, hypernitrosylated, and depleted of calstabin195. We hypothesized that the loss of sarcoglycans are the central upstream event in muscular dystrophies and that an underlying mechanism of the dystrophic phenotype in sarcoglycanopathy should be

similar to that in DMD. Thus we investigated the role of RyR1 dysfunction in β- sarcoglycan-deficient mice (Sgcb−/− mice; an established murine model of LGMD)225.

Chapter 2, Section 2 Results

A hallmark of muscular dystrophies is limb muscle weakness200. EDL muscle from

Sgcb−/− mice displayed reduced specific force relative to wild type (mean force at 70 Hz stimulation ± SEM: Sgcb−/−, 200 ± 20 kNm-1, vs. WT, 440 ± 30 kNm-1; n = 9

(Sgcb−/−), n = 6 (WT); P <0.01 (T-test)), indicating defective force generation that is independent of muscle size.

To determine whether the observed reductions in muscle specific force were associated with remodeling of the RyR1 macromolecular complex, RyR1 were immunoprecipitated and immunoblotted to assay for post-translational modifications. Skeletal muscle RyR1 channels from Sgcb−/− mice exhibited significantly increased phosphorylation, oxidation, and nitrosylation. Moreover, RyR1 from Sgcb−/− muscle were depleted of calstabin1 (Figure 11). Furthermore, Sgcb-/- mice were treated with the 1,4- benzothiazepine derivative, S107, which inhibits calstabin1 depletion from the RyR1 complex, stabilizes the closed state of the RyR1 channel, and improves muscle strength in mdx mice95. Immunoprecipitation and immunoblotting of RyR1 indicated that there was increased calstabin1 bound to RyR1 in the S107 treated Sgcb−/− mice (Figure 11).

Preserved RyR1-calstabin1 interaction is associated with reduced SR Ca2+ leak, improved

Ca2+ release, muscle function, and exercise capacity226. To study SR Ca2+ release, we loaded isolated fast twitch flexor digitorum brevis (FDB) muscle fibers with the fluorescent Ca2+ indicator Fluo-4 AM and electrically stimulated the fibers to produce tetanic contractions. Ca2+ transients were reduced in FDB myocytes from Sgcb−/− mice.

The S107-treated Sgcb−/− displayed increased Ca2+ transients compared to untreated

Sgcb−/− (Figure 12A-B).

We next measured force production in EDL muscles isolated from mice with or without

S107 treatment. There was a significant increase in EDL specific force in the S107- treated Sgcb−/− mice (Figure 13A). A marked feature of skeletal muscle is its susceptibility to fatigue and recovery. EDL muscles from S107-treated and untreated

Sgcb−/− mice were repeatedly stimulated to tetanic contractions. The degree of force reduction during fatigue as well as the recovery was similar in both groups (Figure 13B-

C). However, EDL from S107-treated mice exhibited increased force production prior to the fatigue protocol. Therefore the EDL from S107-treated mice exhibited higher force production throughout the fatigue protocol and would likely sustain higher levels of work in vivo222. To determine whether the improvements in muscle function corresponded to increased exercise capacity, voluntary running performance was recorded in S107-treated and untreated Sgcb−/− mice. S107-treated Sgcb−/− mice ran longer and faster than untreated mice (Figure 14A-B).

Mitochondrial abnormalities have also been described in murine models227 of muscular dystrophy. Accordingly, ultrastructural analysis of EDL muscles from Sgcb−/− mice revealed many fibers with abnormal mitochondrial morphology, such as swelling and loss of cristae structure. However, the sarcomere ultrastructure appeared normal in the

Sgcb−/− muscle fibers (Figure 15A-B). Interestingly, Ca2+ overload leading to mitochondrial dysfunction has been linked to activation of cell death pathways in δ- sarcoglycan deficient mice, and there may be a similar mechanism in Sgcb−/− mice 228.

Chapter 2, Section 2 Summary

Our data indicate that RyR1 in dystrophic muscle are oxidized, cysteine-nitrosylated, phosphorylated, and depleted of calstabin1, resulting in reduced Ca2+ transients, decreased fast twitch muscle force, and impaired exercise capacity. Furthermore, we show that treating β-sarcoglycan-deficient mice with the RyR stabilizing drug, S107, preserves RyR1-calstabin1 binding, increases SR Ca2+ release, fast twitch muscle force, and improves voluntary exercise capacity.

Chapter 2, Section 2 Methods

Animals

Homozygous β-sarcoglycan deficient mice (Strain: B6.129-Sgcbtm1Kcam/1 J; in this article referred to as Sgcb−/−) were obtained from The Jackson Laboratory (Bar Harbor, ME,

USA)225. The Sgcb−/− mice were backcrossed for several generations into C57Bl/6 background and aged-matched C57Bl/6 mice were used as controls. All experiments with

animals were approved by Columbia University’s Institutional Animal Care and Use

Committee.

Voluntary exercise and S107 treatment

At the beginning of each experiment mice were transferred to individual cages equipped with running wheels and exercise was recorded using a data acquisition system

(Respironics). The mice were acclimated to the running wheels for 7 to 9 days and were randomized into two treatment groups. The first group received S107 (25 mg/100 mL) in the drinking water and the second group received water only. S107 (S107-HCl, FW

245.77) was synthesized as previously described72,229. Mice drank ~9 mL/day (water bottle and body weight were recorded to monitor consumption) for a daily dose of S107 of approximately 1.5 mg. There was no difference in daily water consumption between the treatment groups (mean ± SEM: control, 9.9 ± 0.6 mL, S107, 9.3 ± 0.9 mL; n = 5,

P = NS). Mice were sacrificed using CO2 followed by cervical dislocation and muscles were harvested for functional and biochemical analyses. Investigators performing all aspects of the studies were blinded to the treatment groups.

Muscle function

Tyrode solution (in mM: NaCl 121, KCl 5.0, CaCl2 1.8,3 MgCl2 0.5, NaH2PO4 0.4,

NaHCO3 24, EDTA 0.1, glucose 5.5). Muscles were stimulated to contract using an electrical field between two platinum electrodes (Aurora Scientific). At the start of each experiment the muscle length (L0) was adjusted to yield the maximum force. The force- frequency relationships were determined by triggering contraction using incremental stimulation frequencies (EDL: 0.5 ms pulses at 2 to 150 Hz for 350 ms at supra-threshold voltage). The muscles were allowed to rest between every force-frequency stimulation for

>1 min. At the end of the force measurement, the L0 and weight of the muscles were measured and the muscles were snap frozen in liquid N2. To quantify the specific force, the absolute force was normalized to the muscle cross-sectional area, calculated as the muscle weight divided by the length using a muscle density constant of 1.056 kg*m-3230.

Muscle fatigue protocol

After force-frequency measurements, the EDL muscle was fatigued. The fatigue protocol for the EDL muscle consisted of 50 tetanic contractions (70 Hz, 350 ms duration) given at 2-s intervals.

RyR1 immunoprecipitation and immunoblotting

EDLs were isotonically lysed in 0.5 mL of a buffer containing 50 mM Tris–HCl (pH 7.4),

150 mM NaCl, 20 mM NaF, 1.0 mM Na3VO4, and protease inhibitors. An anti-RyR antibody (4 µg 5029 Ab) was used to immunoprecipitate RyR1 from 250 µg of tissue homogenate. The samples were incubated with the antibody in 0.5 mL of a modified

RIPA buffer (50 mM Tris–HCl pH 7.4, 0.9% NaCl, 5.0 mM NaF, 1.0 mM Na3VO4, 1%

Triton-X100, and protease inhibitors) for 1 h at 4°C. The immune complexes were

incubated with protein A Sepharose beads (Sigma, St Louis, MO, USA) at 4°C for 1 h and the beads were washed three times with buffer. Proteins were separated on SDS-

PAGE gels (6% for RyR1, 15% for calstabin1) and transferred onto nitrocellulose membranes for 1 h at 200 mA (SemiDry transfer blot, Bio-Rad). After incubation with blocking solution (LICOR Biosciences, Lincoln, NE, USA) to prevent non-specific antibody binding, immunoblots were developed with anti-RyR (Affinity Bioreagents,

Bolder, CO, USA; 1:2,000), and anti-Cys-NO antibody (Sigma, St Louis, MO, USA;

1:2,000), or an anti-calstabin antibody (1:2,500). To determine channel oxidation the carbonyl groups on the protein side chains were derivatized to 2,4- dinitrophenylhydrazone (DNP-hydrazone) by reaction with 2,4 dinitrophenylhydrazine

(DNPH). The DNP signal on RyR1 was detected by immunoblotting with an anti-DNP antibody. All immunoblots were developed and quantified using the Odyssey Infrared

Imaging System (LICOR Biosystems, Lincoln, NE, USA) and infrared-labeled secondary antibodies.

Ca2+ imaging in FDB muscle fibers

Single FDB fibers were obtained by enzymatic dissociation as previously described222

[30]. FDB muscles from both hind limbs were incubated for approximately 2 h at 37°C in approximately 4 mL Dulbecco’s Modified Eagles Medium (DMEM) containing 0.3% collagenase 1 (Sigma) and 10% fetal bovine serum. The muscles were transferred to a culture dish containing fresh DMEM (approximately 4 mL) and gently triturated until the muscles were dissociated. The cell suspension was stored in an incubator at 37°C/5%

2+ CO2 until the start of the experiment. FDB fibers were loaded with the fluorescent Ca

indicator Fluo-4 AM (5 µM, Invitrogen/Molecular probes) for 15 min in RT. The cells were allowed to attach to a laminin-coated glass cover slip that formed the bottom of a perfusion chamber. The cells were then superfused with tyrode solution (in mM: NaCl

121, KCl 5.0, CaCl2 1.8, MgCl2 0.5, NaH2PO4 0.4, NaHCO3 24, EDTA 0.1, glucose 5.5; bubbled with O2/CO2 (95/5%)). The fibers were triggered to tetanic contraction using electrical field stimulation (pulses of 0.5 ms at supra-threshold voltage, at 70 Hz for 350 ms) and Fluo-4 fluorescence was monitored using confocal microscopy (Zeiss LSM 5

Live, 40x oil immersion lens, excitation wavelength was 488 nm and the emitted fluorescence was recorded between 495 nm and 525 nm) in linescan mode. Only cells that were firmly attached to the glass bottom dish throughout the tetanic stimulation were included in the analysis. After subtraction of background fluorescence, the change in fluorescent signal during the tetanus (peak–resting (ΔF)) was divided by the resting signal (ΔF/F0). All experiments were performed at RT (approximately 20°C). The investigators were blinded to the genotype and treatment of subjects.

Transmission electron microscopy

EDL muscles were fixed in 2.5% glutaraldehyde in 0.1 M Sorenson’s buffer (PH 7.2) followed by 1 h of post-fixation with 1% OsO4 in Sorenson’s buffer. After dehydration the tissue samples were embedded in Lx-112 (Ladd Research Industries) and 60 nm sections were cut using an ultramicrotome (MT-7000). The sections were then stained with uranyl acetate and lead citrate and examined under an electron microscope (JEM-

1200 EXII, JEOL) and images were taken using an ORCA-HR digital camera

(Hamamatsu) and recorded with an AMT Image Capture Engine.

Figure 6: β-adrenergic pathway. (A) Schematic representation of β-adrenergic pathway, beginning with agonist (isoproterenol) binding to β-adrenergic receptor and ending with adenylyl cyclase (AC)-dependent formation of cAMP (step 4), which activates PKA (step 5).

Figure 7: RyR1-S2844A muscle exhibits similar force to WT muscle. (A) Force- frequency curves of extensor digitorum longus (EDL) muscle from WT and RyR1- S2844A. All pooled data are represented as mean±SEM, WT, n=8 muscles; RyR1- S2844A, n=8 muscles.

Figure 8: β-adrenergic inotropy in skeletal muscle is dependent on phosphorylation of serine 2844 in RyR1. (A and C), representative twitch force in extensor digitorum longus (EDL) muscle from (A) WT and (C) 2844A mice. Force was normalized to the maximal twitch force before application of isoproterenol (ISO), which was set to 1. (B and D), average specific force (twitch force normazlied to cross-sectional area) before and after ISO treatment in (B) WT and (D) S2844A EDL muscles. All pooled data are represented as mean±SEM, n=8 muslces for all groups. *P<0.01 (paried t-test for comparisons before and after ISO in the same muscle), n.s., difference between control and ISO in the S2844A group was not significant (paired t-test).

Figure 9: Blocking phosphorylation of RyR1 at S2844 inhibits the β-adrenergic effect on Ca2+ transients in FDB muscle fibers. (A and B), fluorescence signals from cells loaded with the Ca2+ indicator fluo-4 during twitch stimulation before and after ISO application in WT (A) and S2844 (B) flexor digitorum brevis (FDB) muscle fibers. The Fluo-4 signal was normalized to the minimum fluorescence signal before onset of the Ca2+ transient, which was set to zero. (C) Pooled data showing the fold change of the peak Ca2+ transient after application of ISO. Dashed line indicates no change in the presence of ISO. All pooled data are represented as mean±SEM, WT n=8 mice, S2844A n=9 mice. **P<0.01 WT±ISO, ##P<0.01 WT vs. S2844A (paired t-test).

Figure 10: Abolished phosphorylation of RyR1 following β-adrenergic stimulation and reduced in vivo force production in S2844A mice. (A) Representative immunoblot of immunoprecipitated RyR1 from WT and S2844A EDL muscles. The muscles treated with ISO are indicated with +; (B) Forelimb grip strength normalized to body weight in stressed WT and S2944A mice. Mice were allowed to grab on to a metal grid attached to a force transducer with their forelimbs. The maximum grip strength was measured as the mouse was pulled by its tail until it lost its grip. All pooled data are represented as mean±SEM, WT n=8 mice, S2844A n=9 mice. *P<0.05 (unpaired t-test).

Figure 11: RyR1 in β-sarcoglycan deficient muscle is cysteine-nitrosylated, oxidized and depleted of calstabin1. Representative immunoblot of immunoprecipitated RyR1 from WT and Sgcb-/- EDL muscles. The muscle from a WT mouse treated with S107 is marked +; (B) Bar graphs show quantification of immunoblots in (A).

Figure 12: β-sarcoglycan deficient muscle displays defective SR Ca2+ release that is restored by S107 treatment. (A) Representative tetanic Ca2+ transients (normalized Fluo-4 fluorescence) in FDB muscle fibers from WT, β-sarcoglycan-deficient (Sgcb−/−), and S107-treated β-sarcoglycan-deficient (Sgcb−/− S107) mice. (B) Average Ca2+ transient amplitudes. All pooled data are represented as mean±SEM, WT n = 6, Sgcb−/− n =20, Sgcb−/− S107 n = 26 cells from three mice in each group, * P <0.05, ** P <0.01 (ANOVA).

Figure 13: S107 treatment increases muscle force in β-sarcoglycan deficient mice. (A) Force-frequency curves of EDL muscle from WT control, Sgcb−/−, and S107- treated Sgcb−/− mice; (B) Fatigue stimulation (50 tetani; each tetanic stimulation had a duration of 350 ms and was produced by stimulating the muscle with 0.5 ms pulses at 70 Hz frequency) on the same muscles as (A). (C) Relative decline in force production during fatigue in (B). Pooled data are represented as mean ± SEM, n = 6–9 muscles per group, * P <0.05 (ANOVA).

Figure 14: S107 treatment increases voluntary exercise in β-sarcoglycan deficient mice. (A and B) Exercise capacity in Sgcb−/− mice is improved by S107. Daily voluntary running distance (A) and average running speed (B). Pooled data are presented as mean ± SEM, n = 5-8 mice in each group, * P <0.05 (ANOVA).

Figure 15: EDL muscles from β-sarcoglycan deficient mice exhibit abnormal mitochondrial morphology. Representative electron microscopy images of EDL muscle from (A) WT and (B) Sgcb−/− mice. Arrows indicate normal mitochondria, (A) or mitochondria with abnormal morphology, including low cristae density, (B). Images from 11 fibers and two mice in each group were investigated under blinded conditions. The sample is magnified at × 25,000. Scale bar indicates 500 nm.

Chapter 3: Skeletal Muscle Aging: An acquired form of Muscular Dystrophy

Chapter 3, Section 1: Ryanodine receptor oxidation causes muscle weakness in aging.

A hallmark of aging is progressive loss of muscle mass (sarcopenia) and declined skeletal muscle function, characterized by reduced force generating capacity 145,146. Age- dependent deterioration of muscle function is not restricted to mammals as it is also observed in many animals including the nematode, C. elegans 147. Indeed, loss of muscular strength is highly predictive of frailty, disability and mortality with increased age 148-150. Progressive development of muscle dysfunction is common in aged humans and in animal models of aging 150, and affects as many as 30-50% of 80 year olds leading to profound loss of function in the elderly 146.

Much attention has been focused on understanding how to reverse age-related muscle wasting, but there are no established treatments for sarcopenia at this time 150-152. In contrast, improving specific force production, which is also significantly reduced in aged muscle 153,154, has received less attention. The observed loss of specific force in aged muscle suggests that E-C coupling may be impaired in aging. Maladaptive cAMP- dependent PKA-mediated phosphorylation and redox-dependent modifications (cysteine- nitrosylation and oxidation) of the RyR1 have been linked to impaired Ca2+ handling and contractile dysfunction in chronic muscle fatigue, heart failure and muscular dystrophy

95,215,231,232. Defective SR Ca2+ release has been reported in age-dependent muscle

weakness 201,202 and maladaptive modifications of the RyR1 macromolecular complex have been implicated in this condition 203. However, the mechanism underlying impaired

SR Ca2+ release in aging muscle is unknown.

Overall oxidative activity increases in an age-dependent manner in muscle 108,233 and stress-induced protein oxidation increases with age 234,235. Both nitrosylation and oxidation have been shown to affect skeletal muscle RyR1 function and Ca2+ signaling

106,236,237. Jang et al showed that ablation of the antioxidant, SOD1, increased superoxide levels in murine skeletal muscle resulting in reduced specific force and accelerated age- dependent muscle pathology 108. Increased levels of ROS in aged muscle are associated with altered Ca2+ handling. Mice with a malignant hyperthermia mutation (Y522S) in

RyR1 exhibit SR Ca2+ leak, which promotes mitochondrial dysfunction and oxidative stress-mediated changes of RyR1 109. Moreover, oxidation-dependent modifications of

RyR1 can increase SR Ca2+ leak and a vicious cycle between RyR1-mediated SR Ca2+ leak and mitochondrial ROS production has been proposed 109.

In the present study, we explored the role of RyR1 dysfunction as an underlying mechanism for defective Ca2+ handling in age-dependent muscle weakness.

Chapter 3, Section 1 Results:

We examined EDL muscles from aged (24 month old) WT mice to explore the potential role of RyR1 dysfunction in aging skeletal muscle. EDL from aged mice exhibited reduced specific force compared to EDL from young (3-6 month old) controls (Figure

16A-C). Similarly, isolated fast twitch muscle FDB fibers from aged mice exhibited significantly reduced tetanic Ca2+ transients compared to FDB fibers from young mice

(Figure 16D-F).

To determine if the observed reduction in muscle specific force and tetanic Ca2+ release were associated with oxidation-dependent remodeling of the RyR1 macromolecular complex, RyR1 from aged and young adult murine muscles were immunoprecipitated and immunoblotted for components of the RyR1 complex 95. Skeletal muscle RyR1 complexes from aged mice exhibited significantly increased nitrosylation and oxidation, and depletion of calstabin1, compared to channels from young adult mice (Figure 17).

Transient increases in mitochondrial [Ca2+] enhance ATP production 238,239, whereas prolonged and excessively elevated mitochondrial [Ca2+] impair mitochondrial function due to dissipation of the mitochondrial membrane potential (ΔΨm) and increased ROS production 238,239. To test whether RyR1 mediated SR Ca2+ leak can lead to mitochondrial dysfunction we measured changes in mitochondrial/cytosolic [Ca2+] and mitochondrial membrane potential in FDB muscle fibers in the presence of rapamycin (15 µM), which causes SR Ca2+ leak by disrupting RyR1-calstabin1 interactions 86,240. FDB fibers loaded with the fluorescent indicator, Rhod-2, were used to monitor relative mitochondrial

[Ca2+] 241,242. Fibers were first co-stained with Mitotracker Green and Rhod-2 to confirm that Rhod-2 was loaded into mitochondria (Figure 18A-C). Rapamycin treatment caused a time-dependent increase in the Rhod-2 signal, indicating intracellular Ca2+ leak- mediated Ca2+ accumulation in the mitochondria (Figure 18D).

Mitochondrial membrane potential was then measured in FDB fibers using the fluorescent indicator, tetra-metyl rhodamine ethyl ester, TMRE 241,243,244. Rapamycin caused progressive reduction of the TMRE signal (Figure 19A). At the end of each experiment, an uncoupler of the mitochondrial membrane potential (carbonyl cyanide p- trifluoromethoxyphenylhydrazone; FCCP) was added to the muscle fiber resulting in a further reduction in TMRE fluorescence. Rapamycin has other activities in addition to disrupting RyR1-calstabin1 interactions including inhibition of mTOR signaling. The pharmacological agent, FK506, similarly depletes calstabin1 from RyR1 channels in an mTOR-independent manner, as it is involved in the calcineurin pathway 86. Thus treatment with FK506 served as a control for the mTOR-dependent effects of rapamycin treatment. Pharmacological depletion of calstabin1 from RyR1 with FK506 caused reduced mitochondrial membrane potential, similar to rapamycin treatment (Figure 19B).

Together, these results indicate that inhibiting the RyR1-calstabin1 interaction leads to

Ca2+ leak and disruption of mitochondrial membrane potential.

Mitochondrial dysfunction is typically associated with increased ROS production. The fluorescent indicator, MitoSOX Red, was used to measure mitochondrial superoxide

* 245 (O2 ) production . Rapamycin caused an increase in the MitoSOX Red signal (Figure

20A). At the end of each experiment the electron transport chain inhibitor Antimycin A

(10 µM) was applied as a positive control 246. Ca2+ leak and ROS production in muscle are associated with increased reactive nitrogen species (RNS) 109 and increased nitrosylation of the RyR1 was observed in skeletal muscle from aged mice (Figure 17).

We loaded FDB fibers with the fluorescent RNS indicator, DAF-FM (DAF), to examine the effects of leaky RyR1 on RNS production. In the presence of rapamycin, DAF fluorescence increased significantly compared to untreated controls (Figure 20B), consistent with increased RNS production. Taken together, our data indicate that acute induction of RyR1-mediated SR Ca2+ leak leads to defective mitochondrial function associated with elevated ROS and RNS production.

The small molecule, S107, inhibits SR Ca2+ leak by decreasing the stress-induced dissociation of calstabin1 from RyR1 95,226,247. Therefore, we sought to determine whether the rapamycin-induced Ca2+ leak and the consequent detrimental effects on mitochondrial membrane potential would be prevented by treatment with S107. FDB fibers from young

WT mice were incubated with S107 (5 µM) for 2-3 hours before the start of the experiment 248. The rapamycin-induced increase in Rhod-2 fluorescence as well as the loss of mitochondrial membrane potential and increase in MitoSOX Red and DAF signals were prevented by S107 (Figures 18B, 19A, and 20A-B) while intermittent twitch stimulation of the FDB fibers (which causes large but transient increases in cytoplasmic

[Ca2+]) did not alter the mitochondrial membrane potential (Figure 19A). These data indicate that pathologic SR Ca2+ leak, but not action potential-mediated SR Ca2+ release, has detrimental effects on mitochondrial function and that the effects of pathologic Ca2+- leak on mitochondrial membrane potential may be prevented by S107 treatment.

To determine whether inhibiting RyR1 Ca2+ leak improves muscle force and exercise capacity, aged mice were housed individually in cages equipped with running wheels,

and voluntary running time and distance were continuously recorded. Half of the aged mice (n = 13) were supplied with S107 (~50 mg kg-1 d-1) in their drinking water for a 4- week period, while the other half (n = 14) served as the control group. Water consumption was not different between the S107 and vehicle groups (average daily consumption in ml, aged + S107, 8.4 ± 0.63; aged, 7.9 ± 0.4; ± SEM, P = NS). The S107 treated mice exhibited significantly increased running distance (Figure 21A). To determine the mechanism underlying the improved exercise capacity in S107 treated aged mice we measured muscle force production. There was a significant increase in specific force at all stimulation frequencies in the EDL muscles from the S107 treated mice

(Figure 21B-D).

EDL muscles used in the force measurements were also analyzed for post-translational modifications of RyR1. There were no differences in the levels of nitrosylation, oxidation or PKA phosphorylation of RyR1 from muscles from mice treated with S107 vs. control.

This suggests that the oxidative protein modifications are irreversible, as has been reported 249. Protein nitrosylation may have a half-life longer than the 4 weeks that the mice were subjected to S107 treatment in the current study250. Muscle from the S107 treated mice did, however, show significantly more calstabin1 bound to the RyR1 channel complexes compared to those from control mice (Figure 17).

To determine whether the improvement in exercise and skeletal muscle specific force were associated with improved SR Ca2+ release, Ca2+ responses to tetanic stimulation were recorded in FDB myocytes from S107 treated and control aged mice. Tetanic Ca2+

transients were significantly increased in FDB myocytes from S107 treated mice compared to those from the control group (Figure 22A-C).

Chapter 3, Section 1 Summary

In the present study we show that oxidized RyR1 in skeletal muscle from aged mice are depleted of calstabin1, resulting in reduced tetanic Ca2+, decreased muscle specific force and also that aged animals exhibit impaired exercise capacity. Moreover, the small molecule rycal drug, S107, which preserves RyR1-calstabin1 binding and stabilizes

RyR1 channels, improved tetanic Ca2+, restored muscle specific force and improved exercise capacity in aged WT mice. This study is consistent with previous reports showing impaired Ca2+ release in aged muscle 202, reduced SR Ca2+ release in SR vesicles

203 and reduced caffeine-induced release of the SR Ca2+ store 251. In addition to the pathological SR Ca2+ leak via remodeled RyR1 that we report in the present study, other defects such as uncoupling between the voltage sensor and RyR1 202 may also contribute to age-dependent reduction in skeletal muscle specific force.

*- *- Mitochondria are the primary source of superoxide (O2 ) and, in the presence of NO, O2 facilitates the production of reactive nitrogen species and protein nitrosylation 252.

Skeletal muscle RyR1 is sensitive to redox changes 253 and we have previously shown that leaky RyR1 in muscular dystrophy and in muscle fatigue after extreme exercise are cysteine-nitrosylated and depleted of calstabin1. In this study we find that skeletal muscle

RyR1 from aged mice are oxidized and nitrosylated. Furthermore, acute induction of SR

Ca2+ leak causes increased ROS and RNS production. Taken together these data suggest

that SR Ca2+ leak may exacerbate mitochondrial dysfunction by causing mitochondrial

Ca2+ overload, which in turn leads to increased RNS and ROS production. The increase in oxidative stress would further promote RyR1 leak by further oxidizing the channel and depleting it of the stabilizing subunit calstabin1 (Figure 23A-C).

Chapter 3, Section 1 Methods

Exercise wheel experiments and S107 treatment

At the beginning of each experiment the animals were transferred to individual cages equipped with running wheels and exercise patterns were continuously recorded using a data acquisition system from Respironics. The mice were allowed to acclimate to the running wheels for a period of 7-9 days and randomized into two treatment groups. The first group received S107 (50 mg/kg/day) in the drinking water while the second group received water only. S107 (S107-HCl FW 245.77) was synthesized as previously described 229,231,247. Mice drank ~8 ml/day (water bottle and body weight were recorded to monitor consumption) for a daily dose of S107 of ~1.5 mg (~50 mg/kg/day). Mice were sacrificed using excess CO2 followed by cervical dislocation and muscles were harvested for functional and biochemical analyses. Investigators performing all aspects of the studies were blinded to the treatment groups. All experiments with animals were approved by Columbia University’s Institutional Animal Care and Use Committee.

Muscle function

Extensor digitorum longus (EDL) muscles were dissected from hind limbs. Stainless steel hooks were tied to the tendons of the muscles using nylon sutures and the muscles were

mounted between a force transducer (Harvard Apparatus) and an adjustable hook. The muscles were immersed in a stimulation chamber containing O2/CO2 (95/5%) bubbled

Tyrode solution (in mM: NaCl 121, KCl 5.0, CaCl2 1.8, MgCl2 0.5, NaH2PO4 0.4,

NaHCO3 24, EDTA 0.1, glucose 5.5). The muscle was stimulated to contract using an electrical field between two platinum electrodes (Aurora Scientific).

At the start of each experiment the muscle length was adjusted to yield the maximum force. The force–frequency relationships were determined by triggering contraction using incremental stimulation frequencies (EDL: 0.5 ms pulses at 2–150 Hz for 350 ms at supra-threshold voltage). Between stimulations the muscle was allowed to rest for ~1 min. At the end of the force measurement, the length (L0) and weight of the muscle was measured and the muscle was snap frozen in liquid N2. To quantify the specific force, the absolute force was normalized to the muscle cross-sectional area, calculated as the muscle weight divided by the length using a muscle density constant of 1.056 kg*m-3 230.

RyR1 immunoprecipitation and Immunoblotting

EDLs were isotonically lysed in 0.5 ml of a buffer containing 50 mM Tris-HCl (pH 7.4),

RIPA buffer (50 mM Tris-HCl pH 7.4, 0.9% NaCl, 5.0 mM NaF, 1.0 mM Na3VO4, 1%

Triton- X100, and protease inhibitors) for 1 hour at 4°C. The immune complexes were incubated with protein A Sepharose beads (Sigma, St. Louis, MS) at 4°C for 1 hour and

the beads were washed three times with buffer. Proteins were separated on SDS-PAGE gels (6% for RyR1, 15% for calstabin1) and transferred onto nitrocellulose membranes for 1 hour at 200 mA (SemiDry transfer blot, Bio-Rad). After incubation with blocking solution (LICOR Biosciences, Lincoln NE) to prevent non-specific antibody binding, immunoblots were developed with anti-RyR (Affinity Bioreagents, Bolder, CO 1:2,000), anti-phospho-RyR1-pSer2844 (1:5000), an anti-Cys-NO antibody (Sigma, St. Louis, MO,

(DNPH). The DNP signal on RyR1 was determined by immunoblotting with an anti-DNP antibody. All immunoblots were developed and quantified using the Odyssey Infrared

Imaging System (LICOR Biosystems, Lincoln, NE) and infrared-labeled secondary antibodies. For the in vitro oxidation and nitrosylation experiment, skeletal SR membranes (100 µg) were resuspended in 200 µl of buffer (10 mM Tris-HCl, pH 7.2 with complete protease inhibitors (Roche Applied Science)). The samples were treated with 1 mM H2O2 and/or 100 µM NOC-12 (1-Hudroxy-2-oxo-[N-ethyl-2-aminoethyl]-3- ethyl-1-triazen; A.G. Scientific, USA) for 30 min at room temperature. After the reaction was completed, RyR1 was immunoprecipitated from the sample with an RyR antibody and the immunoprecipitates were analyzed for total RyR1, oxidized RyR1, and

Calstabin1 associated with the channel complex as described.

Ca2+ imaging using Fluo-4 AM in FDB muscle fibers

Single FDB fibers were obtained by enzymatic dissociation as previously described 241.

FDB muscles from both hind limbs were incubated for ~2 hours at 37ºC in ~4 ml

Dulbecco’s Modified Eagles Medium (DMEM) containing 0.3% collagenase 1 (Sigma) and 10% fetal bovine serum. The muscles were transferred to a culture dish containing fresh DMEM (~4 ml) and gently triturated using a 1000 µl pipette until the muscles were dissociated. The cell suspension was stored in an incubator at 37ºC/5% CO2 until the start of the experiment.

FDB fibers were loaded with the fluorescent Ca2+ indicator Fluo-4 AM (5µM,

Invitrogen/Molecular probes) for 15 min in RT. The cells were allowed to attach to a laminin coated glass cover slip that formed the bottom of a perfusion chamber. The cells were then superfused with tyrode solution (in mM: NaCl 121, KCl 5.0, CaCl2 1.8, MgCl2

0.5, NaH2PO4 0.4, NaHCO3 24, EDTA 0.1, glucose 5.5; bubbled with O2/CO2 (95/5%)).

The fibers were triggered to tetanic contraction using electrical field stimulation (pulses of 0.5 ms at 70 Hz for a duration of 350 ms, at supra-threshold voltage) and Fluo-4 fluorescence was monitored using a confocal microscope system (Zeiss LSM 5 Live, 40x oil immersion lens, excitation wavelength was 488 nm and the emitted fluorescence was recorded between 495-525 nm). The use of the single excitation/emission dye fluo-4 necessitates normalizing to pre-stimulation values to negate possible differences in dye loading and excitation strength. However, we used similar loading conditions and confocal configurations throughout and we did not observe any differences in baseline fluorescence values among the groups (young: 19.5 AU, (n=8); aged: 17.7 AU (n=20); aged + S107: 16.8 AU (n=16). Furthermore, we examined the fluo-4 accumulation rates

in FDB fibers from young vs. aged mice and found no differences in the rate of dye uptake (young: 5.9 AU/min (n=3); aged 6.3 AU/min (n=3). After subtraction of background fluorescence, the change in fluorescent signal during the tetanus [peak– resting (ΔF)] was divided by the resting signal (ΔF/F0). All experiments were performed in RT (~20ºC). The investigators were blinded to the genotype, age and treatment of subjects.

Measurement of mitochondrial membrane potential

Enzymatically dissociated FDB fibers were incubated with the fluorescent indicator of mitochondrial membrane potential tetra-methylrhodamine ethyl ester (TMRE; 30 nM;

Invitrogen by Life Technologies/Molecular Probes, USA) for 20 minutes at room temperature. Cells were perfused with Tyrode solution (135 mM NaCl, 1 mM MgCl2, 4 mM KCl, 1 mM Glucose, 2 mM HEPES, 1.8 mM CaCl2). In some experiments cells were pre-incubated with S107 (5 µM) for 2-4 hours at 37oC. Using a confocal laser scanning microscope (Zeiss LSM 5 Live, 40x oil immersion lens), TMRE fluorescence was exited at 532 nm laser light and the emitted signal was collected through a band pass filter (540-

625 nm). Fluorescence changes in mitochondria-rich regions was followed over time and quantification was made using ImageJ. In each cell, at each time point, three different mitochondrial-rich regions were analyzed and averaged. Fluorescence measurements were normalized to the baseline fluorescence (Fo) in the same cell at the start of the experiment. Statistical significance was tested using analysis of variance (two-way repeated measures ANOVA).

To test the effect of action potential-induced Ca2+ transients on mitochondrial membrane potential, muscle fibers were repeatedly stimulated with electrical pulses (0.5 ms duration, 1 Hz frequency at supravoltage threshold). To minimize movement artifacts, the muscle fibers were pre-incubated for 30 minutes with N-benzyl-p-toluenesulphonamide

(BTS; 25-125 µM; Toronto Research Chemicals, Canada) 254. Stimulation was discontinued when the fibers were perfused with FCCP.

Ca2+ Imaging using Rhod-2

Enzymatically dissociated FDB fibers were incubated with the fluorescent indicator

Rhod-2/AM (Invitrogen by Life Technologies/Molecular Probes, USA; 5 µM) for ~1 hour at room temperature. To confirm that Rhod-2 was taken up in the mitochondria, cell were simultaneously loaded with the fluorescent indicator Mitotracker green; Invitrogen by Life Technologies/Molecular Probes, USA; 0.2 µM; 30 min room temperature). In mouse FDB fibers, the mitochondria are lined up along the sides of the z-lines and when

Rhod-2 and Mitotracker Green are loaded into the mitochondria this is shown as a striated pattern of fluorescence 241,255. Cells were perfused with Tyrode solution and, when indicated, rapamycin (15 µM) containing Tyrode solution. In some experiments, cells were pre-incubated with S107 (5 µM) for 2-4 hours. Rhod-2 fluorescence was measured using a confocal microscope ((Zeiss LSM 5 Live, 40x oil immersion lens) with excitation at 532 nm and the emitted signal collected through a band pass filter (540-625 nm). Mitotracker green was excited at 488 nm and the emitted fluorescence collected at

>510 nm. ImageJ was used to quantify changes in Rhod-2 fluorescence in mitochondria

rich areas (confirmed by overlapping Rhod-2 and mitotracker green signals). Statistical significance was tested using analysis of variance (ANOVA).

Reactive Nitrogen Species (RNS) Detection

FDB fibers were incubated with 4-amino-5-methylamino-2′,7′-difluorofluorescein diacetate (DAF-FM; 10 µM; Invitrogen by Life Technologies/Molecular Probes, USA), a pH-insensitive fluorescent dye that emits increased fluorescence after reaction with an active intermediate of NO 109, for 20 minutes at room temperature. Cells were perfused with Tyrode solution, incubated with Rapamycin, and perfused with the NO donor S-

Nitroso-N-acetylpenicillamine (SNAP; 100 nM). DAF fluorescence was measured using a confocal microscope (Zeiss LSM 5 Live, 40x oil immersion lens) where the cells were excited at 488 nm and the emitted signal was collected through a band pass filter (495-

525 nm). Images were analyzed using ImageJ.

Mitochondrial Superoxide Detection

FDB fibers were incubated for 20 min at room temperature with MitoSOX Red

(Invitrogen /Molecular Probes, USA; 2.5 µM), a mitochondria-targeted fluorescent indicator for ROS 241. Cells were perfused with Tyrode solution, incubated with

Rapamycin (2.5 µM) and perfused with Antimycin A (10 µM) at the end of the experiment to cause an increase in mitochondrial superoxide production. Using a confocal microscope (Zeiss LSM 5 Live, 40 x oil immersion lens), MitoSOX-loaded cells were excited at 488 nm and the emitted signal was filtered through a band pass filter

(540-625 nm).

Statistics

To determine statistical significance, student’s t-test was used for comparison between two groups. For comparison between more than two groups analysis of variance

(ANOVA) followed by the Bonferroni post hoc test was used. All pooled data are expressed as mean ± SEM unless otherwise noted. All experiments were performed by blinded observers.

Chapter 3, Section 2: Genetically enhancing mitochondrial anti-oxidant activity improves muscle function in aging

Both oxidation of RyR1 and the subsequent intracellular Ca2+ leak underlie the age- dependent reduction in skeletal muscle specific force 220. Acute induction of RyR1- mediated SR Ca2+ leak with rapamycin, which competes the channel-stabilizing subunit, calstabin1 off from RyR1 95,240, resulted in defective mitochondrial function associated with elevated free radical production 220. However the role of mitochondrial ROS in age- dependent reduction in skeletal muscle function has not been elucidated.

Recently there have been numerous efforts to study mitochondria-derived free radicals in health- and lifespan by experimentally expressing catalase, which catalyzes the decomposition of hydrogen peroxide to water and oxygen, in the mitochondria. This has been done using in vitro models 127, adeno-associate viral vectors (AAV) 128 and most recently by genetically engineering its overexpression in mice 129. These transgenic mice,

MCat mice, in which the human catalase is targeted to and overexpressed in mitochondria, display a 10-20% increase in maximum and median lifespan 129, reduced age-related insulin resistance 130, and attenuated energy imbalance. Since mitochondrial targeted overexpression of catalase results in reduced mitochondrial ROS 129,130, here we used the MCat mouse model to investigate the relationship between antioxidant activity and skeletal muscle aging and subsequent functional decline.

Chapter 3, Section 2 Results

6-month-old and 24-month-old MCat and WT littermates were housed individually for 3 weeks in cages equipped with running wheels and voluntary running performance was recorded. Aged MCat mice exhibited significantly increased running distance relative to age-matched WT mice (Figure 24A). This finding correlated with increased time spent on running wheels (Figure 24B).

Since muscle force production is an essential determinant of exercise capacity 148, we hypothesized that mitochondrial ROS contribute to age-dependent reduction in skeletal muscle force generating capacity. To test this we measured force in EDL muscles from young and aged WT and MCat mice. Isolated EDL muscles were electrically stimulated to contract and force production was measured and normalized to cross-sectional area

(yielding a measure of muscle specific force). There were no significant difference in specific force between young WT and MCat muscles. However, EDL muscle from aged

MCat mice exhibited significantly higher specific force than muscles from WT littermates (Figure 25A-D).

Appropriate SR Ca2+ release is essential to skeletal muscle contraction, and we thus studied tetanic Ca2+ transients in enzymatically dissociated FDB muscle fibers loaded with the fluorescent Ca2+ indicator, Fluo-4 AM. Cells were electrically stimulated to produce tetanic contractions and fluorescence was recorded. Ca2+ transients in aged WT and MCat myocytes were markedly reduced relative to young cells. However this age- dependent reduction in Ca2+ transients was significantly improved in aged MCat myocytes (Figure 26A-E). These changes in Ca2+ transients are observable in the absence of a significant difference in resting Ca2+ (Figure 26F).

Skeletal muscle excitation-contraction coupling requires Ca2+ reuptake by the SR Ca2+

ATPase 1 (SERCA1). SERCA1 pumps Ca2+ back into the SR following intracellular Ca2+ release, lowering the cytosolic [Ca2 +] to baseline levels of ~100 nM, thereby causing relaxation (Figure 1A). SERCA1 is tightly regulated by its redox state, and its activity is reduced in aged murine skeletal muscle 256. Thus, we hypothesized that enhanced SERCA activity mechanistically underlies the enhancement of skeletal muscle function in aged

MCat muscle. However, activity of SERCA1 in aged WT skeletal muscle was not significantly different from that in aged MCat littermates (Figure 27A). Furthermore, there was no significant difference in SERCA1 tyrosine nitration in MCat vs. age- matched WT littermates (Figure 27B). Overall SERCA1 expression in WT vs. MCat littermates was consistent as well (Figure 27C).

We and others have shown that SR Ca2+ leak is associated with impaired exercise

capacity, defective Ca2+ handling, and dysfunctional skeletal muscle performance 72,257.

To test the hypothesis that RyR1-mediated SR Ca2+ leak is decreased in aged MCat mice, we measured Ca2+ sparks in permeabilized FDB muscles 258. We found a significant reduction in Ca2+ spark frequency in aged MCat muscles compared with WT littermates

(Figure 28A-B). Depletion of the SR Ca2+ store is a consequence of increased SR Ca2+ leak in aged skeletal muscle 251. Therefore, we hypothesized that reducing oxidative stress by genetically enhancing mitochondrial catalase activity would prevent this Ca2+ depletion in MCat mice. Though SR Ca2+ load was reduced in aged WT and MCat relative to their young counterparts, aged MCat muscle exhibited significantly higher SR

Ca2+ load than aged WT (Figure 28C).

Preserved RyR1-calstabin1 interaction is linked to reduced SR Ca2+ leak 95,220.

Furthermore, RyR1 oxidation and cysteine nitrosylation decrease the binding affinity of calstabin1 for RyR1 106,259, eventually resulting in leaky channels associated with intracellular Ca2+ leak and increased Ca2+ sparks. Oxidation-dependent post-translational modifications of RyR1 affect skeletal muscle force generating capacity and this is a key mechanism in age-dependent muscle weakness 220. We therefore examined whether age- dependent oxidative remodeling of the RyR1 macromolecular complex is reduced in

MCat mice. RyR1 from aged and young EDL muscles were immunoprecipitated and immunoblotted for components of the RyR1 complex and concomitant redox modifications 95,220. Age-dependent RyR1 oxidation and cysteine-nitrosylation were both reduced in MCat skeletal muscle, and there was more calstabin1 associated with channels from aged mutant animals compared to WT littermates (Figure 29A). Overall expression

of neither RyR1 nor calstabin1 was altered in aged WT relative to aged MCat muscles

(Figure 27C).

To assess the single channel properties of RyR1 in its remodeled state, SR membranes were prepared from EDL muscles and fused to planar lipid membrane bilayers, and Ca2+

86,220 fluxes through RyR1 channels were recorded . The open probability (Po) of skeletal muscle RyR1 channels from young mice was low, as expected for normal skeletal muscle

RyR1 channels (Figure 29B-C). In contrast, skeletal muscle RyR1 channels from aged

WT mice exhibited a significantly increased Po relative to those from aged MCat mice

(Figure 29B-C).

Chapter 3, Section 2 Summary

In the present study we used a genetic model with enhanced mitochondrial antioxidant activity (MCat mice) to investigate the effects of increased mitochondrial antioxidative capacity on age-dependent loss of skeletal muscle function and Ca2+ signaling. Our data indicate that MCat mice exhibit reduced age-dependent loss of muscle function. Aged

MCat mice exhibited improved voluntary exercise, increased skeletal muscle specific force and tetanic Ca2+ transients, decreased intracellular Ca2+ leak and increased sarcoplasmic reticulum Ca2+ load when compared with age-matched WT littermates.

Furthermore, RyR1 from aged MCat mice were less oxidized, had more calstabin1 associated with it, and exhibited increased single channel open probability. We thus provide compelling evidence for a direct role of mitochondrial free radicals in promoting the pathological intracellular Ca2+ leak that underlies age-dependent loss of skeletal

muscle function. This study harbors implications for the development of novel therapeutic strategies, including mitochondria-targeted antioxidants for treatment of mitochondrial myopathies.

Chapter 3, Section 2 Methods

Animals

MCat mouse model generation has been described previously 129. MCat founders were kindly provided by Peter S. Rabinovitch (University of Washington, Seattle, WA) and backcrossed >9 generations onto C57Bl/6 background. We used 6-month old (young) and

24-month old (aged) male MCat mice and their age-matched WT littermates. The rodents were housed in a 22°C room with a 12-hour light/dark cycle and were allowed food and water ad libitum. Blinded observers performed all experiments.

Voluntary exercise

Analysis of voluntary exercise was performed using individual cages equipped with running wheels. To minimize environmental differences, mice were allowed to acclimate to the running wheels for a period of at least 7 days and then exercise on the wheel was continuously recorded for 2 weeks using a data acquisition system (Minimitter, Bend,

OR). Mice were euthanized and muscles were quickly harvested for functional and biochemical analyses.

Muscle function

The specific force of whole extensor digitorum longus (EDL) muscles was assessed ex

vivo using a system from Aurora Scientific (Ontario, Canada). EDL muscles were dissected and stainless steel hooks were tied to the tendons of the muscles using nylon sutures and the muscles were mounted between a force transducer (Harvard Apparatus,

Holliston, MA) and an adjustable hook. The muscles were immersed in a stimulation chamber containing O2/CO2 (95/5%) bubbled Tyrode solution (in mM: NaCl 121, KCl

5.0, CaCl2 1.8, MgCl2 0.5, NaH2PO4 0.4, NaHCO3 24, EDTA 0.1, glucose 5.5). The muscle was stimulated to contract using an electrical field between two platinum electrodes. At the start of each experiment the muscle length was adjusted to yield the maximum force. The force–frequency relationships were determined by triggering contraction using incremental stimulation frequencies (EDL: 0.5 ms pulses at 2–150 Hz for 350 ms at supra-threshold voltage). Between stimulations the muscle was allowed to rest for ~1 min. The fatigue protocol consisted of 50 tetanic contractions (70 Hz, 350 ms duration) given at 2-s intervals. At the end of the force measurement, the length and weight of the muscle was measured and the muscle was snap frozen in liquid nitrogen. To quantify the specific force, the absolute force was normalized to the muscle cross- sectional area, calculated as the muscle weight divided by the length using a muscle density constant of 1.056 kg*m-3.

Ca2+ transients measurement

Single flexor digitorum brevis (FDB) fibers were obtained by enzymatic dissociation.

FDB muscles from both hind limbs were incubated for ∼2 h at 37°C in ∼4 ml Dulbecco's

Modified Eagles medium containing 0.3% collagenase 1 (Sigma, St Louis, MO, USA) and 10% fetal bovine serum. The muscles were transferred to a culture dish containing

fresh Dulbecco's modified Eagle's medium (∼4 ml) and gently triturated until the muscles were dissociated. The cell suspension was stored in an incubator at 37°C/5% CO2 until the start of the experiment.

FDB fibers were loaded with the fluorescent Ca2+ indicator fluo-4 acetoxymethyl ester

(AM; 5 µM, Invitrogen/Molecular Probes, Grand Island, NY, USA) for 15 min at room temperature. The cells were allowed to attach to a laminin-coated glass coverslip that formed the bottom of a perfusion chamber. The cells were then superfused with Tyrode solution [in mM: NaCl 121, KCl 5.0, CaCl2 1.8, MgCl2 0.5, NaH2PO4 0.4, NaHCO3 24,

EDTA 0.1, glucose 5.5; bubbled with O2/CO2 (95/5%)]. The fibers were triggered to twitch contraction using electrical field stimulation (pulses of 0.5 ms at supra-threshold voltage) and fluo-4 fluorescence was monitored using a confocal microscope system

(Zeiss LSM 5 Live, 40× oil immersion lens, excitation wavelength was 488 nm and the emitted fluorescence was recorded between 495 and 525 nm). The use of single excitation/emission dye Fluo-4 necessitates normalizing to pre-stimulation values to negate possible differences in dye loading and excitation strength. Only fibers attached to the bottom of the perfusion chamber throughout the twitch stimulation were measured from.

SERCA1 activity assay

SERCA activity was measured using the malachite green procedure for phosphate determination, adapted to the microscale as previously described 260. The reaction was started by the addition of 50 µg of muscle microsomes to 150 µl of reaction mixture (20

mM MOPS/Tris-HCl, pH 6.8. 100 mM KCl, 5 mM MgCl2, 5 mM ATP, 1 mM EGTA.

2+ 0.350 mM CaCl2 (free Ca concentration of approximately 500 nM as calculated using the CHELATOR program). After 5 min, the reaction was stopped by the transfer of 120

µl of reaction mixture to 80 µl of malachite green reagent mixture in a 96-well microplate. The malachite green reagent mixture was made by mixing 0.122% malachite green hydrochloride in 6.2 N H2SO4, 5.76% ammonium paramolybdate tetrahydrate, and

11% Tween 20 in a volume ratio of 100:66:2. Color development was quenched after

10 s by the addition of 45 µl of 15.1% sodium citrate dihydrate. Inorganic phosphate liberated in the ATPase reaction was quantified by comparison of absorbance at 570 nm with standard curves generated with known amounts of Na2HPO4 in the reaction buffer.

Ca2+ sparks measurement

For Ca2+ sparks measurements, FDB muscle fibers were permeabilized in a relaxing solution (in mM: 140 K-glutamate, 10 HEPES, 10 MgCl2, 0.1 EGTA, pH 7.0) containing

0.01% saponin for ~1 min. After washing the sample with a saponin free solution, the solution was changed to an internal medium (in mM: 140 potassium L-glutamate, 5

Na2ATP, 10 glucose, 10 HEPES, 4.4 MgCl2, 1.1 EGTA, 0.3 CaCl2 (free [Ca2+] 130 nM), Fluo-3 0.05; pentapotassium salt, Invitrogen, USA; pH 7.0). Fluorescence images were acquired with a Zeiss LSM 5 Live confocal system (63× oil immersion, NA=1.4) operated in line-scan mode (x vs. t, 2 ms/line, 5000 lines /scan) along the longitudinal axis of the fibers. Fluo-3 was excited with an Argon laser at 488 nm, and the emitted fluorescence was recorded between 495-555 nm. Ca2+ sparks detection and analyses used custom made routines compiled in IDL (v7.1, ITT) according to algorithms described

previously 261,262.

Immunoprecipitation and Immunoblot Analysis

Muscle samples (EDL) were isotonically lysed in 0.5 ml of buffer containing 50 mM

Tris–HCl (pH 7.4), 150 mM NaCl, 20 mM NaF, 1.0 mM Na3VO4 and protease inhibitors.

The samples were incubated with the antibody in 0.5 ml of a modified RIPA buffer (50 mM Tris–HCl pH 7.4, 0.9% NaCl, 5.0 mM NaF, 1.0 mM Na3VO4, 1% Triton-X100 and protease inhibitors) for 1 h at 4°C. The immune complexes were incubated with protein A

Sepharose beads (Sigma) at 4°C for 1 h and the beads were washed three times with buffer. Proteins were separated on SDS–PAGE gels (6% for RyR1, 15% for calstabin1,

10% for SERCA1) and transferred on to nitrocellulose membranes for 2 h at 200 mA

(SemiDry transfer blot, Bio-Rad, Hercules, CA, USA). Immunoblots were developed using the following primary antibodies: anti-RyR1 (1:2000, Thermo Scientific, Rockford,

IL), anti-phospho-RyR1-pSer2844 (1:5,000), anti-calstabin (FKBP12 C-19, 1:1000, Santa

Cruz Biotechnology, Inc., Santa Cruz, CA), anti-Cys-NO (1:1000, Sigma-Aldrich), anti-

SERCA1 (1:1000, Abcam, San Francisco, CA), anti-n-tyrosine (1:1000, Abcam, San

Francisco, CA). To determine channel oxidation, the carbonyl groups in the protein side chains within the immunoprecipitate were derivatized to 2,4-dinitrophenylhydrazone

(DNP) by reaction with 2,4-dinitrophenylhydrazine. The DNP signal associated with RyR was determined using a specific anti-DNP antibody, according to the manufacturer’s instructions (Millipore, Billerica, MA). Levels of RyR1 bound proteins were normalized to the total RyR1 immunoprecipitated (arbitrary units). All immunoblots were developed with the Odyssey system (LI-COR Biosciences, Lincoln, NE), using IR-labeled anti-

mouse and anti-rabbit IgG (1:10000 dilution) secondary antibodies, as described 263.

Single Channel Recordings

Muscles were homogenized using a tissue homogenizer (Fisher Scientific) at the highest speed for 1 min with 2 volumes of 20 mM Tris-maleate (pH 7.4), 1 mM EDTA and protease inhibitors (Roche). Homogenate was centrifuged at 4,000 g for 15 min at 4°C and the supernatant was centrifuged at 40,000Xg for 30 min at 4°C. The final pellet, containing the SR fractions, was resuspended and aliquoted in: 250 mM sucrose, 10 mM

MOPS (pH 7.4), 1 mM EDTA and protease inhibitors. Samples were frozen in liquid nitrogen and stored at -80°C. SR vesicles containing RyR1 were fused to planar lipid bilayers formed by painting a lipid mixture of phosphatidylethanolamine and phosphatidylcholine (Avanti Polar Lipids) in a 3:1 ratio across a 200 µm hole in polysulfonate cups (Warner Instruments) separating 2 chambers. The trans chamber (1.0 ml), representing the intra-SR (luminal) compartment, was connected to the head stage input of a bilayer voltage clamp amplifier. The cis chamber (1.0 ml), representing the cytoplasmic compartment, was held at virtual ground. Symmetrical solutions used were as follows (in mM): 1 mM EGTA, 250/125 mM Hepes/Tris, 50 mM KCl, 0.64 mM

CaCl2, pH 7.35 as cis solution and 53 mM Ca(OH)2, 50 mM KCl, 250 mM Hepes, pH

7.35 as trans solution. The concentration of free Ca2+ in the cis chamber was calculated with WinMaxC program (version 2.50; www.stanford.edu/~cpatton/maxc.html). SR vesicles were added to the cis side and fusion with the lipid bilayer was induced by making the cis side hyperosmotic by the addition of 500 mM KCl. After the appearance of potassium and chloride channels, the cis side was perfused with the cis solution.

Single-channel currents were recorded at 0 mV by using a Bilayer Clamp BC-525C

(Warner Instruments), filtered at 1 kHz using a Low-Pass Bessel Filter 8 Pole (Warner

Instruments), and digitized at 4 kHz. All experiments were performed at room temperature (23°C). Data acquisition was performed by using Digidata 1322A and

Axoscope 10.1 software (Axon Instruments). The recordings were analyzed by using

Clampfit 10.1 (Molecular Devices) and Origin software (ver. 6.0, Microcal Software).

SR Ca2+ load determination in isolated FDB muscle cells

FDB muscle cells were preloaded with 5 µM low affinity Ca2+ dye mag-fluo-4 for 30 minutes, then 1 mM 4-CmC was applied to cells to induce maximum release. Mag-fluo-4 was excited at 488 nm and emission was collected at 495-525.

Resting [Ca2+] determination

Resting [Ca2+] in FDB muscle cells were ratiometrically measured with fluo-4 and fura- red as described previously 264. FDB muscle cells were simultaneously loaded with 5 µM fluo-4 AM and 10 µM fura-red for 10 min. They were then excited at 488 nm and emission was collected at 495-525 (F515) and 650-700 (F675), respectively. The ratio of the two emissions represents [Ca2+].

Statistics

In all the experiments mice were coded to ‘blind’ investigators with respect to genotype.

The sample size (‘n’ in each group) for each experiment is stated in the figure legends.

Data are expressed as mean ± standard error (SEM), unless otherwise indicated. To

determine statistical significance we used two-way ANOVA and comparison t test, as appropriate. Bonferroni post hoc testing was performed where applicable. Minimum statistically significant differences were established at p < 0.05.

Figure 16: Impaired force production and reduced SR Ca2+ release in EDL muscle from aged mice. (A and B) Tetanic contractions of EDL muscle from young (A) and aged (B) mice (force normalized to cross-sectional area); (C) Average force at the indicated stimulation frequencies in EDL muscles from young and aged mice (mean, ± SEM, n = 5 (young), 7 (aged), P < 0.05 among groups at all stimulation frequencies); (D and E) Normalized Fluo-4 fluorescence in FDB muscle fibers during a 70 Hz tetanic stimulation; (F) Peak Ca2+ responses in FDB fibers stimulated at 70 Hz (fibers taken from the same animals as in A and B; mean, ± SEM, n = 8 (young), 10 (aged), * P < 0.05).

Figure 17: RyR1 from aged muscle is cysteine-nitrosylated, oxidized and depleted of calstabin1. Representative immunoblot of immunoprecipitated RyR1 from young and aged mice. The muscle from a WT mouse treated with S107 is marked. DNP: 2,4- dinitrophenylhydrazone. P*RyR1: Phosphorylated RyR1 (at serine 2844). Cys NO: cysteine nitrosylation; (B) Bar graphs show quantification of immunoblots in (A).

Figure 18: Pharmacological dissociation of RyR1 and calstabin1 causes increased mitochondrial Ca2+. (A-C) Mitochondrial uptake of the Ca2+ indicator Rhod-2 AM in FDB muscle fiber. Image showing an FDB muscle fiber loaded with the fluorescent indicators Rhod-2 (A) and Mitotracker Green (B). In murine FDB fibers, the mitochondria are lined up along the sides of the z-lines and when the fluorescent indicators are loaded into the mitochondria a striated pattern is seen. (C) Overlap between the Ca2+ indicator, Rhod-2 and the mitochondrial marker, Mitotracker Green. The scale bar indicates 10 µM; (D) Rapamycin-induced increase in mitochondrial Ca2+ measured with the fluorescent indicator, Rhod-2. The Rhod-2 signal was measured from three mitochondria rich regions in each cell (mitochondria rich regions were confirmed using mitotracker green, A-C) and normalized to baseline; * P < 0.05 indicates significant difference of the control rapamycin (N = 7) compared to S107 (N = 6) and control no rapamycin (N = 4) groups (ANOVA). In the S107 rapamycin group FDB fibers were incubated with S107 (5 µM) for 2-4 hrs before starting the experiment.

Figure 19: Pharmacological dissociation of RyR1 and calstabin1 causes reduced mitochondrial membrane potential. (A) Changes in mitochondrial membrane potential (measured with TMRE fluorescence) with respect to different interventions. Arrow indicates onset of rapamycin (for groups control rapamycin and S107 rapamycin) or repetitive twitch stimulation without rapamycin (twitching). The dashed line indicates application of FCCP (300 nM). * P < 0.05 indicates significant difference (ANOVA) among the groups control rapamycin (n = 5) and control no rapamycin (n = 3) or group twitching (n = 6) or S107 rapamycin (n = 5); (B) FK506-induced decrease in mitochondrial membrane potential. Arrow indicates onset of FK506 (50 µM) treatment. The dashed line indicates application of FCCP (300 nM). Data is shown as mean ± SEM. * P < 0.05 indicates significant difference (ANOVA) between the two groups control no FK506 (n = 4) and FK506 (n = 4).

Figure 20: Pharmacological dissociation of RyR1 and calstabin1 causes increased mitochondrial ROS and RNS production. (A) Mitochondrial superoxide production in FDB fibers measured with MitoSOX Red. Arrow indicates when rapamycin was applied. The dashed line indicates application of Antimycin A (10 µM) as a positive control for superoxide production. Control rapamycin (n = 8), Control no rapamycin (n = 5), S107 rapamycin (n = 5). * P < 0.05 indicates significant difference between the control rapamycin group and no rapamycin or S107 groups (ANOVA); (B) The effect of rapamycin-induced Ca2+ leak on RNS production in FDB fibers measured with the RNS indicator DAF. Arrow indicates application of rapamycin. The NO donor S-nitroso-N- acetylpenicillamine (SNAP; 100 nM) was applied as a positive control at the end of each experiment (indicated by dashed line). Control rapamycin (n = 6), control no rapamycin (n = 6), S107 rapamycin (n = 5). * P < 0.05 indicates significant difference (ANOVA) for the control rapamycin group compared to the S107 and no rapamycin groups. All data are shown as mean ± SEM.

Figure 21: Improved exercise capacity and muscle specific force following S107 treatment of aged mice. (A) Daily voluntary running distance in aged mice ± S107 treatment (mean, ± SEM, n = 13 aged + S107, n = 14 aged, * P < 0.05, ANOVA). The arrow indicates start of the S107 treatment; (B and C) 70 Hz tetanic contractions in isolated EDL muscles from aged and S107 treated aged mice; (D) Average specific force in EDL muscles from the same mice as in A (mean ±SEM, n = 6 young, 7 aged, * P < 0.05, *** P < 0.001).

Figure 22: Increased SR Ca2+ release following S107 treatment of aged mice. (A and B) Ca2+ transients Fluo-4 fluorescence in FDB muscle fibers during a 70 Hz tetanic stimulation in control mice (A) and mice treated with S107 (B); (C) Peak tetanic Ca2+ amplitudes in the two treatment groups (muscle fibers were taken from the same animals as in Figure 3A-B; mean, ±SEM, n = 10-13, ** P < 0.01).

Figure 23: Model of RyR1-mediated SR Ca2+ leak and mitochondrial dysfunction in aging skeletal muscle. (A) Sarcoplasmic reticulum (SR) Ca2+ leak due to oxidation- dependent modifications of RyR1 exacerbates mitochondrial dysfunction and production of reactive oxygen species (ROS). This causes remodeling of RyR1 resulting in SR Ca2+ leak, which impairs muscle force production. (B-C) The RyR1 from young mice are not “leaky”, the SR Ca2+ stores are filled and activation of the myocyte leads to SR Ca2+ release, which triggers muscle contraction. (D-E) In aging, ROS and reactive nitrogen species (RNS)–mediated remodeling of RyR1 results in dissociation of the RyR1 stabilizing subunit calstabin1 and SR Ca2+ leak. Under these conditions, muscle activation will lead to reduced SR Ca2+ release and impaired muscle force. Ryanodine receptor type 1: RyR1 Reactive oxygen species: ROS. Sarcoplasmic reticulum: SR. 2+ 2+ Mitochondrial [Ca ]: [Ca ]m. Mitochondrial membrane potential: ΔΨm.

Figure 24: Improved exercise capacity in aged MCat mice. Mice were housed in individual cages equipped with running wheels for three weeks. Exercise distance (A) and running wheel time (B) were recorded. Data are mean ± SEM (*: p<0.01 vs. young WT; #p<0.05 vs. aged WT; n: young WT=7, young MCat=8, aged WT=8, aged MCat=8, ANOVA).

Figure 25: Preserved skeletal muscle function in aged MCat mice. (A and B) Tetanic contractions (70 Hz) in isolated EDL muscles from MCat and WT littermates (force normalized to cross-sectional area); (C and D) Average specific force in EDL muscles from the same mice as in panels (A) and (B). Data are mean ± SEM (n: young WT=4, young MCat=4, aged WT=8; aged MCat=7; t-test was performed for each individual point: ∗: p<0.05 vs. aged WT).

Figure 26: Improved tetanic Ca2+ in skeletal muscle from aged MCat mice. (A-D) Representative traces of normalized Fluo-4 fluorescence in FDB muscle fibers during a 70 Hz tetanic stimulation; (E) Peak Ca2+ responses in FDB fibers stimulated at 70 Hz (fibers taken from the same animals as in A-D, n = 15-21 cells from at least 3 mice in each group); (F) Resting cytosolic Ca2+ (measured ratiometrically). Data are mean ± SEM (∗: p<0.05 vs. young WT; #: p<0.05 vs. aged WT, ANOVA).

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Figure 27: SERCA1 activity in MCat mice is similar to WT. (A) SERCA1 activity in skeletal muscle; (B) Representative immunoblot from experiments of immunoprecipitated SERCA1 from murine skeletal muscle; (C) Bar graphs show quantification of immunoblots in (B); (D) Representative immunoblot using skeletal muscle lysates using anti-RyR1, anti-SERCA1, anti-Calstabin1 and anti-Tubulin (loading control) antibodies (*: p<0.01 vs. young WT, #: p<0.01 vs. young MCat, ANOVA); (E) Bar graphs show quantification of immunoblots in (D).

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Figure 28: Reduced SR Ca2+ spark activity and increased SR Ca2+ load in muscle from aged MCat mice. (A) Representative images of line scans of Fluo-4 fluorescence from permeabilized FDB muscle fibers showing Ca2+ spark activity. The heat diagram indicates the normalized change in fluorescence intensity (ΔF/F0); (B) Bar graph showing average Ca2+ spark frequency (n=15-25 cells from at least 3 mice in each group); (C) Representative time course of Ca2+ leak from SR microsomes following Ca2+ uptake; (D) Ca2+ leak as calculated by the percentage of uptake; (E) SR Ca2+ load (measured by applying 1 mM 4-CmC). Data are mean ± SEM (*: p<0.05, **: p<0.01 vs. young WT; #: p<0.05 vs. aged WT, ANOVA).

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Figure 29: Skeletal muscle RyR1 isolated from aged MCat mice is biochemically remodeled and exhibits reduced single-channel open probability (Po). (A) Representative immunoblots of immunoprecipitated RyR1 from aged murine EDL. DNP: 2,4-dinitrophenylhydrazone; (B) Representative RyR1 single-channel current traces. Channel openings are shown as upward deflections and the closed (c-) state of the channel is indicated by horizontal bars in the beginning of each trace. Tracings from over 2 min of recording for each condition showing channel activity at two time scales (5 s in upper trace and 500 ms in lower trace) as indicated by dimension bars, and the respective Po (open probability), To (average open time), and Tc (average closed time) are shown above each trace. The activity of the channel indicated by the thick black bar is shown on the expanded time scale (the 500 ms trace below); (C) Bar graph summarizing Po at 150 nM cytosolic [Ca2+] in young WT (n=6), aged WT (n=5), young MCat (n=7), and aged MCat (n=5) channels. Data are mean ± SEM (*: p<0.05, **: p<0.01 vs. young WT, #: p<0.05, ## p<0.01 vs. aged WT, ANOVA).

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Chapter 4: The role of the UNC-68/intracellular calcium release channel in age- dependent loss of muscle function in C. elegans

Strikingly, despite an approximately 2,000-fold difference in the lifespans of humans and

C. elegans204, both exhibit age-dependent reductions in muscle function and motor activity that ultimately contribute to senescence and death. Due to its short lifespan and well-characterized genome, C. elegans has been used as a model for the study of the genetics of aging and lifespan determination 157,162 including the age-dependent decline in locomotion147,205,206. Age-dependent reduction in locomotion in C. elegans has been attributed to degeneration of the nervous system 207 and the body wall musculature 147 208, however the molecular determinants of age-dependent muscle deterioration and declining motility have not been determined.

unc-68 is the RyR gene in the C. elegans genome 176. unc-68(e540) null mutants exhibit locomotive defects characterized by the “unc”, or “uncoordinated” phenotype156.

Treatment with the RyR-binding drug, ryanodine, induces contractile paralysis in wild- type C. elegans, but unc-68 (e540) are unaffected by ryanodine 156,176,178. Ca2+ transients triggered by action potentials in C. elegans body wall muscles require UNC-68173,178. The intracellular Ca2+ release channel shares 40% homology with human RyR1 179 (Figure 5A), and important functional domains are conserved including the PKA phosphorylation site and two EF- hand motifs180. The C. elegans genome contains eight FKBs with varying levels of sequence identity compared to mammalian calstabins (Figure 5B). The one with the

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highest homology to a mammalian isoform is fkb-2, with ~60% sequence identity to its mammalian homolog, calstabin181. We hypothesized that UNC-68 is composed of a regulatory macromolecular signaling complex including FKB-2. Furthermore, we investigated the role of UNC-68 throughout nematode lifespan.

Chapter 4 Results

We hypothesized that a macromolecular complex similar to that of mammalian RyRs regulates UNC-68. To test this hypothesis, lysates were prepared from populations of freeze-cracked wild type (WT) C. elegans and UNC-68 was immunoprecipitated using our mammalian anti-RyR antibody (5029). The immunoprecipitates were immunoblotted to detect UNC-68, the catalytic subunit of protein kinase A (PKAcat), protein phosphatase

1 (PP1), FKB-2, and phosphodiesterase 4 (PDE-4), using mammalian anti-RyR, anti-

PKA, anti-PP1, and anti-calstabin antibodies. The previously published C. elegans anti-

PDE-4265 was used to detect PDE-4 on the channel. UNC-68 comprises a macromolecular complex similar to that found in the mammalian muscle that includes

PKAcat, PP1, PDE-4 and FKB-2 (Figure 30A-B). FKB-2 was depleted from UNC-68 in the fkb-2 (ok3007) putative null mutant (Figure 30A).

A band that immunoprecipitated with the anti-calstabin (FKBP12) antibody was not part of the UNC-68 macromolecular complex (Figure 30B) in the experiments conducted for completion of this dissertation. Presence of this band in fkb-2(ok3007) was confirmed through Western blot analysis (Supplementary Figure 1). Since the antibody used for the immunoprecipitation experiment was not suitable for immunoblotting FKB-2 in

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nematode samples (data not shown), wild type and fkb-2(ok3007) samples were immunoblotted against FKB-2 using an alternate anti-Calstabin1 antibody

(Supplementary Figure 1). Immunoprecipiation experiments were also conducted to confirm that this band is not associated with the UNC-68 macromolecular complex in fkb-2(ok3007). Subsequent experiments in the Marks laboratory did not yield this band

(F. Forrester, personal communication, February 2015), thus it will be important to further characterize fkb-2(ok3007). Specifically, the standard in the field would be to demonstrate that the effects that the mutant confers are due to the lesion by criteria such as rescue, silencing of the gene with RNAi, or an independent allele.

However, it is important to note that neither of the anti-Calstabin1 antibodies used detected a band in fkb-2(ok3007) when the samples were immunoprecipitated for UNC-

68 and immunoblotted against Calstabin1, demonstrating that FKB-2 is missing from the

UNC-68 macromolecular complex in this mutant. Thus, this mutant is well-suited for the experiments presented in this dissertation.

Since the UNC-68 macromolecular complex was intact in this model organism, we investigated age-dependent regulation of the channel. RyR1 channels are oxidized, leaky and Ca2+ transients are reduced in aged mammalian skeletal muscle220. Ca2+ transients were measured in partially immobilized transgenic nematodes expressing the genetically encoded Ca2+ indicator, Pmyo-3::GCaMP2, in the body wall muscle cells173,266 (Figure

31A-B). Wild type C. elegans exhibited an age-dependent decline in body wall muscle peak Ca2+ transients from day 3 to day 15 post-hatching (Figure 31C).

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Furthermore, RyR1 oxidation has been linked to SR Ca2+ leak and impaired muscle function during extreme exercise and in heart failure and muscular dystrophies 72,81,95.

Our group has reported that oxidation of RyR1 and the subsequent intracellular Ca2+ leak are underlying mechanisms of age-related loss of skeletal muscle specific force (force normalized to the cross sectional area of muscle) 220. UNC-68 was depleted of FKB-2, and oxidized (Figure 32A) in an age-dependent manner 267. The FKB-2 depletion from the UNC-68 complex occurred despite changes in overall FKB-2 expression throughout nematode lifespan (Figure 32B).

Genetic FKB-2 deficiency caused acceleration of the age-dependent reduction in body wall muscle Ca2+ transients (Figure 31C) with peak Ca2+ in fkb-2(ok3007) worms at day 7 being significantly lower than that of wild type of the same age. Similarly, UNC-68 was significantly more oxidized at day 7 in fkb-2 mutants compared to WT (Figure 32A).

To investigate the effect of FKB-2 dissociation from UNC-68 in aged worms, independent of the other confounding variables inherent to aging, pharmacological interventions mimicking the aged state in young adult nematodes were applied. FKB-2 was competed off from the UNC-68 macromolecular complex using rapamycin or

FK506. Both rapamycin and FK-506 bind to calstabin and compete it off from RyR channels resulting in leaky channels and release of SR Ca2+ in the resting state 86,267,268.

Rapamycin-FKBP12 inhibits mTOR and FK506 inhibits calcineurin. Therefore their effects downstream of those related to RyR are unrelated. It is thus likely that their non-

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RyR related actions do not account for the observed SR Ca2+ leak.

Age-synchronized young C. elegans were treated with rapamycin or FK506. Acute treatment with FK506 and rapamycin each independently caused depletion of FKB-2 from the channel (Figure 33). Furthermore, longer treatment with FK506 caused oxidation of UNC-68, demonstrating a relationship between depletion of FKB-2 and oxidation of UNC-68 (Figure 33).

Ca2+ transients were measured in partially immobilized transgenic nematodes, Pmyo-

3::GCaMP2173,266. Pharmacologic depletion of FKB-2 from UNC-68 by rapamcyin or

FK506 treatment caused reduced body wall muscle Ca2+ transients in WT (Figure 34A).

When FKB-2 was genetically depleted from the UNC-68 complex as in the fkb-2 mutants, treatment with rapamycin or FK506 had no effect on the Ca2+ transients (Figure

4B). These data suggest that rendering UNC-68 channels leaky by removing FKB-2 depletes SR Ca2+ resulting in reduced Ca2+ transients and weakened muscle contraction.

To investigate the individual effect of age-dependent UNC-68 oxidation independent of the other confounding variables involved in aging147, we introduced another pharmacological intervention mimicking the aged state in young adult nematodes.

Treating with the superoxide-generating agent, paraquat269 caused increased oxidation of

UNC-68 and depletion of FKB-2 from the channel in a concentration-dependent manner

(Figure 35A). Furthermore, contraction-associated Ca2+ transients decreased with paraquat treatment in a concentration-dependent manner (Figure 35B). These data

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indicate that both UNC-68 oxidation and FKB-2 depletion each independently contribute to the observed aging body wall muscle phenotype.

The small molecule rycal, S107 inhibits SR Ca2+ leak by reducing the stress-induced depletion of calstabin from the RyR channel complex95,247. Here, we show that treatment with S107 (10µM) for 3-5 hours re-associated FKB-2 with UNC-68 in aged nematodes

(Figure 36A). Furthermore, treatment with S107 improved peak Ca2+ in an FKB-2 dependent manner (Figure 36B-C).

Chapter 4 Summary

In the present chapter we show that UNC-68 is comprised of a macromolecular complex with striking similarities to that of RyR1, including the channel-stabilizing subunit, FKB-

2. Like calstabin, FKB-2 regulates UNC-68 by directly associating with the channel.

Competing FKB-2 from UNC-68 with rapamycin or FK506 results in reduced body wall muscle Ca2+ transients due to intracellular Ca2+ leak. Moreover, as we previously reported in aged mice220, there is age-dependent oxidation of UNC-68, depletion of FKB-2 from the UNC-68 channel complex and reduced Ca2+ transients in aged nematodes. This aging phenotype was accelerated in fkb-2 mutants. Finally, re-associating FKB-2 with UNC-68 in aged C. elegans using the RyR-stabilizing drug, S107, improved Ca2+ transients. Our data provides evidence for age-dependent UNC-68 dysfunction-mediated reduction in body wall muscle Ca2+ transients, and progressive oxidation of UNC-68 over the course of nematode lifespan, which renders the channel leaky.

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Chapter 4 Methods

C. elegans strains and culture conditions

Worms were grown and maintained on standard nematode growth medium (NGM) plates with a layer of OP50 Escherichia coli in an incubator at 20oC, as described156. N2

(Bristol) and fkb-2(ok3007) were provided by the Caenorhabditis Genetics Center

(University of Minnesota). fkb-2(ok3007) was backcrossed six times. The transgenic strain expressing Pmyo-3::GCaMP2 was kindly provided by Zhao-Wen Wang, University of Connecticut Health Center173. Pmyo-3::GCaMP2 was subsequently crossed into fkb-

2(ok3007) for measurement of contraction-associated Ca2+ transients.

Age synchronization

Adult worms at the egg-laying stage were treated with alkaline hypochlorite solution to obtain age-synchronized populations and eggs were plated on NGM plates, as described270. For experiments requiring aged worms, age-synchronized animals at the L4 stage were collected in M9 buffer and plated on NGM plates containing 5-fluoro-2’- deoxyuridine (FUDR, Sigma, 50 µM) to prevent egg-laying271.

Immunoprecipitation and Western Blotting

Nematodes were grown under standard conditions. For protein biochemistry experiments, a procedure to crack nematodes in a solubilizing and denaturing buffer was adapted272.

Briefly, worms were washed and collected with M9 buffer. They were centrifuged for 2 minutes at 1,000 rpm three times to wash. Worms were allowed to settle to the bottom of the collection tube by sitting on ice for ~5 minutes. Fluid was removed and the worm

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pellet was snap frozen with liquid nitrogen. Frozen pellets containing whole nematodes were then rapidly thawed by running the tubes under warm water. A volume of

Nematode solubilization buffer equal to the volume of the worm pellet was added

(Nematode solubilization buffer: 0.3% Ethanolamine, 2 mM EDTA, 1 mM PMSF in

DMSO, 5 mM DTT, 1x protease inhibitor) and tubes were microwaved on medium (25 s for 100 µl pellet; time was increased for greater volumes). Lysates were then quickly drawn into a syringe through a 26-gauge needle and forced back through the needle into a new collection tube on ice. Samples were centrifuged at 1,000 rpm for 2 minutes to remove insoluble material and the supernatant was transferred to a new tube on ice.

Lysates were snap frozen and stored in at -80oC.

A mammalian anti-RyR antibody prepared in our laboratory (4 µg 5029 Ab) was used to immunoprecipitate UNC-68 from 100 µg of nematode homogenate. Samples were incubated with the antibody in 0.5 ml of a modified RIPA buffer (50 mM Tris-HCl pH

7.4, 0.9% NaCl, 5.0 mM NaF, 1.0 mM Na3VO4, 1% Triton- X100, and protease inhibitors) for 1 hour at 4°C. The immune complexes were incubated with protein A

Sepharose beads (Sigma, St. Louis, MS) at 4°C for 1 hour and the beads were washed three times with buffer. Proteins were size-fractionated by SDS-PAGE (6% for UNC-68,

15% for FKB-2) and transferred onto nitrocellulose membranes for 1 hour at 200 mA

(SemiDry transfer blot, Bio-Rad). After incubation with blocking solution (LICOR

Biosciences, Lincoln NE) to prevent non-specific antibody binding, immunoblots were developed using antibodies against RyR (5029, 1:5000), PKAcat (Santa Cruz

Biotechnology, sc-903, 1:1000), PDE4 (kindly provided to us by Kenneth Miller,

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Oklahoma Medical Research Foundation, Oklahoma City, Oklahoma), PP1 (sc6104,

1:1000) or an anti-calstabin antibody (Abcam, ab108420 1:2,500). To determine channel oxidation the carbonyl groups on the protein side chains were derivatized to 2,4- dinitrophenylhydrazone (DNP-hydrazone) by reaction with 2,4-dinitrophenylhydrazine

(DNPH) according to manufacturers (Millipore) instructions. The DNP signal on immunoprecipitated UNC-68 was determined by immunoblotting with an anti-DNP antibody (Millipore, 1:1000). All immunoblots were developed and quantified using the

Odyssey Infrared Imaging System (LICOR Biosystems, Lincoln, NE) and infrared- labeled secondary antibodies. In addition, immunoblotting and immunoprecipitation of the UNC-68 macromolecular complex were conducted using another anti-calstabin antibody (1:2000, Abcam) and the same methods as described.

Imaging contraction-associated body wall muscle Ca2+ transients

Spontaneous changes in body wall muscle Ca2+ were measured in nematodes expressing

GCaMP2 by imaging fluorescence intensity using a Zeiss Axio Observer inverted microscope with an electron-multiplying CCD camera (Photometrics Evolve 512) and an

LED light source (Colibri). Nematodes were partially immobilized by placing them individually into a 5-10 µl drop of M9 buffer, suspended between a glass slide and coverslip. Twenty-second videos of individual nematodes were recorded.

Analyzing contraction-associated body wall muscle Ca2+ transients

Contraction-associated body wall muscle Ca2+ transients were analyzed using an

Interactive Data Language (IDL)-based image quantification software that was developed

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for this purpose in our laboratory. For each twenty-second video, signals from the body wall muscles in nematodes expressing GCaMP2 fluorescence were analyzed using an edge-detection algorithm from each frame as "line-scan" images with the nematode perimeter on the y-axis and time (s) on the x-axis. These images are then quantified based on the average of the peak Ca2+ fluorescence signal on the worm muscle wall (Fig. 3).

Drug treatments

To pharmacologically deplete FKB-2 from UNC-68 nematodes were treated for 15 minutes with 15 µM and 50 µM rapamycin and FK506, respectively. To re-associate

FKB-2 and UNC-68 in aged nematodes, treatment was with 10 µM S107 for 3-5 hours.

Nematodes were grown in standard conditions, age synchronized as described, washed and collected with M9 buffer, centrifuged for 2 minutes at 1,000 rpm three times to wash.

Worms were allowed to settle to the bottom of the collection tube by sitting on ice for ~5 minutes. Fluid was removed and the worm pellet was gently resuspended in M9 containing the appropriate drug concentration and gently rocked on a shaker at room temperature for the denoted time periods. Collection tubes were centrifuged for 2 minutes at 1,000 rpm and M9 containing drug was removed and replaced with M9. Biochemistry or Ca2+ measurements were conducted, as described.

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Figure 30: UNC-68 is comprised of a macromolecular complex. UNC-68 (A) and FKB-2 (B), respectively, were immunoprecipitated and immunoblotted using anti-RyR, anti-phosphodiesterase 4 (PDE4), anti-protein kinase A (catalytic subunit; PKAcat), anti- protein phosphatase 1 (PP1), and anti-calstabin (Santa Cruz) antibodies in murine skeletal sarcoplasmic reticulum preparations (Sk SR), wild type populations of C. elegans (WT) and populations of fkb-2(ok3007); (C and D) Bar graphs show quantification of immunoblots in (A and B).

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Figure 31: Age-dependent reduction in Ca2+ transients is accelerated in fkb-2 relative to WT. (A) Image of representative Pmyo-3::GCaMP2 nematode; top arrow indicates high fluorescence in the contracting muscle, bottom arrow represents low fluorescence in relaxing muscle; (B) processed image used for analysis of Ca2+ levels; (C) Ca2+ transients in age-synchronized populations of wild type and fkb-2 (ok3007) nematodes (***: P<0.001; n=15-30 nematodes for each group; ANOVA). (D) UNC-68 was immunoprecipitated from age-synchronized populations of mutant and WT nematodes and immunoblotted using anti-RyR, anti-Calstabin (Santa Cruz) and DNP (marker of oxidation) antibodies.

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Figure 32: Age-dependent UNC-68 remodeling is accelerated in fkb-2 relative to WT. (A) UNC-68 was immunoprecipitated from age-synchronized populations of mutant and WT nematodes and immunoblotted using anti-RyR, anti-Calstabin (Santa Cruz) and DNP (marker of oxidation) antibodies; (B) Bar graphs show quantification of immunoblots in (A); (C) Nematode lysates were immunoblotted using anti-Calstabin and anti-tubulin (loading control) antibodies; (D) Bar graphs show quantification of immunoblots in (C).

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Figure 33: Depleting FKB-2 from UNC-68 causes UNC-68 oxidation. (A) UNC-68 was immunoprecipitated and immunoblotted using anti-RyR, anti-Calstabin (Santa Cruz) and DNP (marker of oxidation) antibodies in nematodes acutely treated with 15 µM and 50 µM rapamycin and FK506, respectively; (B) Bar graphs show quantification of immunoblots in (A).

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Figure 34: FKB-2 depletion from UNC-68 causes defective Ca2+ handling. Contraction-associated Ca2+ transients measured in young age-synchronized WT (A) and fkb-2(ok3007) (B) populations treated for 15-20 minutes with 15 µM and 50 µM rapamycin (Rap) and FK-506, respectively (**: P<0.01; ***: P<0.001; n=15-30 nematodes for each group; ANOVA).

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Figure 35: UNC-68 oxidation causes defective Ca2+ handling. (A) UNC-68 was immunoprecipitated and immunoblotted using anti-RyR, anti-Calstabin (Santa Cruz) and DNP (marker of oxidation) antibodies in nematodes acutely treated for 15-20 minutes with paraquat; (B) Bar graphs show quantification of immunoblots in (A); (C) Contraction-associated Ca2+ transients measured in young age-synchronized WT nematodes treated for 15-20 minutes with paraquat (*: P<0.05; **: P<0.01 n=15-30 nematodes for each group; ANOVA).

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Figure 36: Treating aged nematodes with the RyR-stabilizing drug, S107, increases body wall muscle Ca2+ transients in aged C. elegans. (A) UNC-68 was immunoprecipitated and immunoblotted with anti-RyR, anti-Calstabin (Santa Cruz) and DNP (marker of oxidation) in aged nematodes with S107 treatment as indicated; (B) Bar graphs show quantification of immunoblots in (A); Contraction-associated Ca2+ transients were measured in age-synchronized WT (C) and (D) fkb-2 mutants treated with S107 as indicated (*: P<0.05; n=15-30 nematodes for each group; ANOVA).

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Supplementary Figure 1: Confirmation that FKB-2 is part of the UNC-68 macromolecular complex. (A) FKB-2 was immunoprecipitated and immunoblotted with anti-RyR and anti-Calstabin (Abcam) in murine skeletal muscle (Sk SR), wild type populations of C. elegans (WT) and populations of fkb-2(ok3007); (B) Murine skeletal muscle (Sk SR), wild type populations of C. elegans (WT) and populations of fkb- 2(ok3007) were immunoblotted using anti-Calstabin antibody (Abcam).

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DISCUSSION

The present study implicates RyR1 dysfunction as important in age-related loss of skeletal muscle function. Acute sympathetic nervous activation increases RyR1 function in muscle contraction. On the other hand chronic stress-induced RyR1 leak contributes to muscle weakness in muscular dystrophies and aging. Our data that suggest that aging is an acquired form of muscular dystrophy, based on the mechanistic similarities underlying the two pathological states. Namely, we find that RyR1 oxidation underlies age- dependent skeletal muscle weakness, and that a source of this oxidation is mitochondria- produced ROS. Finally we demonstrate the remarkable evolutionary conservation of

RyR1 by investigating the regulation of its C. elegans homolog UNC-68.

Post-translational modification of the ryanodine receptor type 1 underlies skeletal muscle weakness in the physiological and pathophysiological state

Chapter 2 is an investigation of the common mechanisms underlying altered skeletal muscle function in the physiological state (Section 1; inotropy) and in the pathophysiological state (Section 2; LGMD). Interestingly, similar post-translational modifications of RyR1 that underlie increased skeletal muscle force in the physiological inotrophy state underlie weakened muscle function in LGMD.

Our laboratory has previously reported that chronic stress induces remodeling of the

RyR1 macromolecular complex such that components of the complex including the phosphodiesterase PDE4D3 (due to oxidation of the channel) and calstabin1 are depleted

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from the RyR1 leading to diastolic SR Ca2+ leak 75,213,221,248,273. These changes, combined with other complex maladaptations that occur under prolonged pathological stress, could contribute to defective muscle function. Here, we investigated the mechanism whereby acute adrenergic stress causes enhanced skeletal muscle function and how this well established physiological process could be explained by PKA phosphorylation of RyR1 at Ser2844. The data show that brief exposure of WT muscles to adrenergic stress leads to transient RyR1-S2844 PKA phosphorylation, providing activation of RyR1 and skeletal muscle contraction downstream of β-adrenergic receptor activation.

Though numerous components of the EC coupling machinery have been excluded as underlying the adrenergic stimulation-mediated increase in skeletal muscle force production including the L-type Ca2+ channel, SR Ca2+ load and myofilament Ca2+ sensitivity 209,216-218,274, the role of RyR1 in adrenergic stimulation-mediated increase of skeletal muscle force production had not been definitively addressed prior to this study.

To resolve this gap we generated a genetic mouse model in which the PKA phosphorylation site in RyR1 was ablated by replacement of serine 2844 with alanine.

Using this new genetic model, we show here that the stress-induced inotropic response in skeletal muscle is dependent on increased SR Ca2+ release via RyR1, and that PKA phosphorylation of a single amino-acid of the RyR1, serine 2844, is the key signal. Thus, we have identified the β-receptor downstream target responsible for increased SR Ca2+ release and skeletal muscle force during sympathetic stimulation as occurs with exercise.

These findings elucidate a fundamental mechanism regulating muscle function. We thus

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hypothesized that similar mechanisms underlie skeletal muscle force enhancement underlie skeletal muscle weakness in the pathophysiological state (Section 2).

In Section 2 we show that remodeling of the RyR1 contributes to skeletal muscle weakness and reduced exercise capacity in Sgcb−/− mice, a model of LGMD. This is consistent with results from a previous study of the mdx mouse model for DMD, in which

RyR1 were S-nitrosylated, and displayed SR Ca2+ leak through RyR195. A null mutation in one of the sarcoglycan proteins results in loss of the whole sarcoglycan complex but not of dystrophin199. However mutations in dystrophin, which cause DMD, also lead to loss of the sarcoglycans200. Thus our findings that RyR1 dysfunction is a common underlying mechanism of DMD and LGMD suggest that the loss of sarcoglycans is a central upstream event in DGC-associated muscular dystrophies.

Cardiomyopathy is a common symptom of muscle dystrophy275 and improved cardiac function has been observed following S107 treatment of heart failure (post-myocardial infarction) and in mdx mice276,277. However the cardiac function was normal as measured by echocardiography in Sgcb-/- mice (data not shown). Sgcb−/− mice that were treated with S107 displayed improved exercise capacity, measured as voluntary running distance and speed. Exercise capacity is a compound measure that involves the function of several organ systems. Therefore, though it is possible that the improved cardiac function in

Sgcb−/− mice following S107 treatment contributed to the improved running capacity, it is likely that muscle function was an underlying mechanism. Muscle function is a central determinant of exercise capacity81 and the reduced tetanic Ca2+ and impaired muscle

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specific force in Sgcb−/− were improved by fixing the skeletal muscle SR Ca2+ leak with

S107.

Skeletal Muscle Aging: An acquired form of Muscular Dystrophy

Taken together, the studies described in Chapter 3 identify mitochondrial ROS-mediated

RyR1 oxidation as an underlying mechanism in age-dependent skeletal muscle dysfunction.

In Section 1 we present data that demonstrates that oxidized RyR1 in muscle from aged mice are depleted of calstabin1, resulting in reduced tetanic Ca2+ release, decreased muscle specific force and impaired exercise capacity. Moreover, S107, which preserves

RyR1-calstabin1 binding and stabilizes RyR1 channels, improved tetanic Ca2+ release, restored muscle specific force and improved exercise capacity in aged WT mice.

Furthermore, acute induction of SR Ca2+ leak with rapamycin or FK506 increased mitochondrial ROS and overall RNS production.

This study is consistent with previous reports showing impaired Ca2+ release in aged muscle 202, reduced SR Ca2+ release in SR vesicles 203 and reduced caffeine-induced release of the SR Ca2+ store 251. It should be noted that S107 treatment did not reduce age-dependent RyR1 oxidation or cysteine-nitrosylation. Oxidation-dependent carbonyl modifications of proteins have been reported to be irreversible 249. Furthermore, the half- life of the mitochondria is 2-4 weeks 278,279 and the half-life of RyR1 is ~10 days in muscle isolated from aged rats 280. Thus, the continued ROS production from

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mitochondria and the slow turnover of the RyR1 protein contribute to the observation that

RyR1 oxidation is not reduced during the four-week course of S107 treatment described in Section 1, despite inhibition of the intracellular Ca2+ leak. Inhibiting SR Ca2+ leak would decrease additional RyR1 oxidation but would not necessarily reverse the oxidation of the Ca2+ release channel. Thus, the investigation of a genetic model with enhanced antioxidative capacity was an important next step in identifying the specific role and source of ROS underlying RyR1 oxidation in aged muscle.

In Section 2, we used a genetic model with enhanced mitochondria-targeted antioxidant activity to investigate the effects of mitochondrial ROS on age-dependent loss of skeletal muscle function and Ca2+ signaling. Indeed, mitochondria-targeted overexpression of catalase improved both whole organism (exercise capacity), and skeletal muscle (specific force) performance, and prevented age-dependent reduction in Ca2+ transients, reduced age-related biochemical modifications of the SR Ca2+ release channel (channel oxidation/nitrosylation and dissociation of calstabin1), and decreased SR Ca2+ leak. We thus provide compelling evidence for a direct role of mitochondrial free radicals in promoting the pathological intracellular Ca2+ leak that underlies age-dependent loss of skeletal muscle function. The results of the two sections presented in Chapter 3 are consistent with a “vicious cycle” whereby SR Ca2+ leak and mitochondrial ROS are locally amplified and lead to progressive age-dependent skeletal muscle dysfunction109

(Figure 23).

Several groups have reported that RyR is tightly regulated by post-translational

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modifications involving remodeling of the ryanodine receptor macromolecular complex

106,109,253,259. Intriguingly, here we show that not only age-dependent RyR1 oxidation, but also cysteine nitrosylation is reduced in MCat mice, and that RyR1 oxidation is accompanied by nitrosylation in aged mice. This finding is consistent with reports that uncovered the capacity of reactive nitrogen species to regulate catalase activity in skeletal muscle 119,121. Thus, catalase overexpression may down-regulate cellular levels of nitroxide free radicals, thereby impacting cysteine nitrosylation of RyR1. The redox- specific post-translational modifications that were attenuated in aged MCat mice were consistent with reduced RyR1-mediated SR Ca2+ leak. This is in agreement with studies in which prolonged exposure to NO donors has been shown to increase SR Ca2+ leak and resting cytosolic Ca2+ in voltage-clamped mouse FDB fibers 281. Additionally, inhibiting

RyR1-mediated SR Ca2+ leak results in rescue of age-dependent increase in spontaneous release of SR Ca2+ (Ca2+ sparks) in permeabilized FDB muscle fibers, as shown in aged

MCat muscle fibers in Section 2.

The regulatory mechanisms of aging are likely multifactorial and several signaling pathways have been reported to contribute, e.g. changes in insulin/insulin-like growth factor (IGF), sirtuin, AMP kinase and inflammatory signaling 150,162. Moreover, mitochondrial dysfunction is strongly implicated in the aging mechanism 282,283. In

Chapter 3, we explore the interplay between RyR1-mediated SR Ca2+ leak, oxidative stress and mitochondrial dysfunction in age-dependent skeletal muscle weakness.

Interestingly, the underlying mechanism of age-dependent skeletal muscle weakness is

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similar to that found in Limb Girdle Muscular Dystrophy (Chapter 2), and aging may thus be characterized as an “acquired” form of muscular dystrophy.

C. elegans: A model for the physiological study of muscle function in aging

Taken together the data presented in Chapter 4 show for the first time that the C. elegans intracellular Ca2+ release channel, UNC-68, comprises a macromolecular complex and its regulation is highly conserved throughout evolution from nematodes to humans. Binding of the mammalian homolog of the stabilizing subunit, calstabin/FKB-2, to the UNC-68 channel is required to prevent a pathological leak of intracellular Ca2+, as is the case in mammalian muscle220. Aged C. elegans have reduced Ca2+ transients, and their UNC-68 channels are oxidized and depleted of FKB-2. Each of these post-translational modifications of UNC-68 likely contribute to the aging phenotype, as pharmacologically depleting UNC-68 of FKB-2 and oxidizing the channel in young nematodes each independently yield the observed aging phenotype of reduced contraction-associated Ca2+ transients. Genetic FKB-2 deficiency causes an accelerated aging phenotype; Ca2+ transients are reduced in younger populations of fkb-2 nematodes and UNC-68 is oxidized at an earlier time point in the lifespan of these mutants relative to WT. Treating aged WT nematodes with the RyR-stabilizing drug, S107, re-associates FKB-2 with

UNC-68 and increases the Ca2+ transients, indicating that UNC-68 dysfunction is likely an underlying mechanism of age-dependent decrease in Ca2+ transients in C. elegans body wall muscle.

We present here for the first time an association between age-dependent oxidation of

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UNC-68 and reduction in Ca2+ transients in aged nematodes. Though the oxidative stress theory of aging was first proposed in 1956284,285, there is still substantial controversy surrounding the role of ROS in aging. For example, deletion or overexpression of the

ROS detoxification enzyme superoxide dismutase (SOD) has little effect on life span in

C. elegans286,287. However, loss of the gene, sesn-1, which encodes sestrin, an evolutionarily conserved protein that is required for regenerating hyperoxidized forms of peroxiredoxins and for ROS clearance, causes reduced lifespan288. Furthermore, ROS levels measured in vivo in C. elegans increase with age 289.

It would be interesting to know whether the increased UNC-68 oxidation and subsequent reduction in body wall muscle Ca2+ transients are a result of globally increased ROS levels or increased ROS levels in UNC-68-surrounding microdomains. For example, we have previously shown that inducing RyR leak in enzymatically dissociated skeletal muscle cells causes increased mitochondrial membrane potential and mitochondrial ROS production220. Based on these data we have proposed a model in which RyR1 leak, due to age-dependent oxidation of the channel, causes mitochondrial Ca2+ overload resulting in

ROS production and further oxidation of RyR1. This further exacerbates the SR Ca2+ leak and creates a vicious cycle between RyR1 and mitochondria that contributes to age- dependent loss of muscle function267. C. elegans would be an excellent model in which to test this model further in aged organisms in vivo.

We also show for the first time that in the putative null mutant, fkb-2(ok3007), FKB-2 does not co-immunoprecipitate with UNC-68. Moreover, the skeletal muscle aging

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phenotype that we characterize in WT nematodes (biochemically modified UNC-68 and reduced Ca2+ transients) is accelerated in fkb-2 mutants. There are eight FKBs that are homologous to mammalian calstabin in the C. elegans genome; FKB-1 and FKB-8 both have ~50% sequence identity to calstabin. Further studies may elucidate the possibility that in the absence of FKB-2 another FKB may stabilize UNC-68. However this appears not to be the case since the aging phenotype is accelerated in the absence of FKB-2.

Another key question is why UNC-68 becomes oxidized within two weeks where the same post-translational modification requires two years in mice and 80 years in humans204.

Taken together, our data indicate that the C. elegans homolog of RyR, UNC-68 comprises a macromolecular complex and is regulated by FKB-2. We have identified an age-dependent reduction in body wall muscle Ca2+ transients in nematodes that is coupled to oxidation and remodeling of UNC-68. Furthermore, our data strongly suggest a role for FKB-2 and UNC-68 in the age-dependent changes in Ca2+ signaling, as treatment with the pharmacological ryanodine receptor stabilizer, S107, increases body wall muscle

Ca2+ transients.

Overall, the data presented in this dissertation demonstrate the evolutionarily conserved mechanisms of skeletal muscle ryanodine receptor regulation in the physiological and pathophysiological states. Acute post-translational modifications of RyR1 channels are pivotal to healthy muscle contraction, and chronic stress-induced post-translational modifications result in leaky RyR1 channels that contribute to skeletal muscle

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dysfunction in age-dependent skeletal muscle weakness and Muscular Dystrophies.

Furthermore, the same post-translational modifications of the ryanodine receptor channel are found in mammals and C. elegans. This work has broad implications for understanding the regulation of skeletal muscle contraction, and important disorders that greatly affect human health. Furthermore, this works presents ryanodine receptor channels as a viable therapeutic target for age-related skeletal muscle weakness,

Muscular Dystrophies, and also implicates C. elegans as a potential model system in which to test future therapeutic targets.

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