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The effects of RYR2 gene deletion on cardiac function and metabolism Bround, Michael J. 2016

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EFFECTS OF RYR2 GENE DELETION ON CARDIAC FUNCTION AND METABOLISM by  Michael J Bround  B.Sc., The University of British Columbia, 2010  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Cell and Developmental Biology)   THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2016  © Michael J Bround, 2016   ii Abstract  The cardiac ryanodine receptor 2 (RYR2) is a sarcoplasmic reticulum Ca2+ release channel central to cardiomyocyte biology. RYR2 Ca2+ release has a well-established role in activating cardiomyocyte motor proteins during excitation-contraction coupling and is therefore critical for heart function. RYR2 is also poised to have other important cardiac functions such as setting heart rate, stimulating ATP metabolism, regulating cardiac hypertrophy, and controlling cardiomyocyte survival. In addition, there is evidence that RYR2 dysfunction occurs during heart disease, suggesting that RYR2 may be a driver of cardiac pathology. The research in this thesis seeks to test which aspects of cardiomyocyte biology are regulated by RYR2 signaling and whether RYR2 loss-of-function is pathogenic. Using a heart-specific, inducible gene deletion system in mice we were able to show that loss of Ryr2 caused heart failure and reduced cardiac contraction. In addition, we saw that Ryr2 deletion lead to reduced heart rate, tachycardic arrhythmia, diminished oxidative metabolism, increased cardiac hypertrophy, and increased cell death via a novel mechanism. To test whether the metabolic and heart rate effects persist in the absence of heart failure, we used an inducible, heart-specific 50% Ryr2 deletion model. In this context we did not see heart failure or decreased cardiac function, but still observed a decrease in heart rate and altered oxidative metabolism. Unlike complete Ryr2 knockout, the 50% Ryr2 ablation model did not display a general decrease in oxidative ATP metabolism, but instead a specific decrease in glucose oxidation. This was associated with reduced mitochondrial Ca2+ uptake and decreased activation of the pyruvate dehydrogenase complex, a Ca2+ sensitive gatekeeper of glucose oxidation. Collectively, these results provide compelling evidence that RYR2 is an essential component of excitation-contraction coupling and a critical driver of   iii cardiac pacemaking. These results also demonstrate that RYR2 is critical for mitochondrial Ca2+ uptake and stimulating oxidative metabolism and strongly suggest that RYR2 has a specific role in activating glucose oxidation. This research also shows that loss of Ryr2 recapitulates heart failure and suggests RYR2 may be involved in hypertrophy and cell death. This suggests a model where RYR2 simultaneously regulates a several facets of cardiomyocyte biology.    iv Preface  I, Michael J Bround, contributed to the research presented in this thesis at all levels. Along with my supervisor, James D Johnson, I conceived the research program presented here. I was substantially involved in the planning and design of the work. I performed a variety of experiments (with the exclusion of those itemized below), and was involved in the analysis of virtually all data presented. I substantially contributed to the interpretation of the data and the conclusions presented here are my own.  The research presented in this thesis is the fruit of collaboration and as such I am indebted to the teamwork of a variety of people whose specific contributions are outlined below.  A version of Chapter 3 has been published. Bround MJ, Asghari P, Wambolt RB, Bohunek L, Smits C, Philit M, Kieffer TJ, Lakatta EG, Boheler KR, Moore ED, Allard MF, Johnson JD. 2012. Cardiac ryanodine receptors control heart rate and rhythmicity in adult mice. Cardiovascular Research, 96:372-80 (PMID: 22869620) Role: Along with my supervisor JD Johnson, I conceived and interpreted all experiments, as well as wrote and edited the manuscript. I also conducted and analyzed the majority of experiments.  Collaborator contributions: P Asghari: Performed EM Spectroscopy experiments and analysis. RB Wambolt: Performed working heart perfusion experiments and analysis. L Bohnuek: Performed echocardiography experiments and analysis.   v C Smits: Coordinated animal work at The James Hogg Research Centre at St. Paul's Hospital M Philit: Performed ECG Telemetry implantation surgeries. TJ Kieffer, EG Lakatta, KR Boheler, ED Moore, and MF Allard contributed key conceptual expertise, experimental resources, and editorial assistance.  A version of Chapter 4 has been published. Bround MJ, Wambolt RB, Luciani DS, Kulpa JE, Rodrigues B, Brownsey RW, Allard MF, Johnson JD. 2013. Cardiomyocyte ATP production, metabolic flexibility, and survival require calcium flux through cardiac ryanodine receptors in vivo. The Journal of Biological Chemistry, 288:18975-86 (PMID: 23678000). Role: Along with my supervisor JD Johnson, I conceived and interpreted all experiments, as well as wrote and edited the manuscript. I also conducted and analyzed the majority of experiments.  Collaborator contributions:  RB Wambolt: Performed working heart perfusion experiments and analysis. JE Kulpa: Performed ATP HPLC measurements and analysis. DS Luciani, B Rodrigues, RW Brownsey, and MF Allard contributed key conceptual expertise, experimental resources, and editorial assistance.  Chapter 5 is based on a manuscript in progress. Bround MJ, Wambolt R, Albu R, Han J, McAfee D, Pourrier M, Scott NE, Bohunek L, Kulpa JE, Fedida D, Brownsey RW, Borchers CH, Foster LJ, Mayor T, Allard MF, Johnson JD. Partial Ryr2 loss disrupts mitochondrial Ca2+ cycling and specifically inhibits glucose oxidation in adult mouse cardiomyocytes. Submitted to JBC in November 2015. Role: Along with my supervisor, I conceived and interpreted all experiments, as   vi well as wrote and edited the manuscript. I also conducted and analyzed the majority of experiments.  Collaborator contributions: R Wambolt: Performed working heart perfusion experiments and analysis. R Albu: Performed protein MS experiments and analysis J Han: Performed metabolomics MS experiments and analysis D McAfee and M Pourrier: Performed cardiomycoyte contraction measurements and analysis. NE Scott: Performed characterization of SILAM heart tissues, assisted with MS analysis. L Bohnuek: Performed echocardiography experiments and analysis. JE Kulpa: Performed ATP HPLC measurements and analysis. D Fedida, RW Brownsey, CH Borchers, LJ Foster, T Mayor, and MF Allard contributed key conceptual expertise, experimental resources, and editorial assistance.     vii Table of Contents  Abstract .......................................................................................................................................... ii	  Preface ........................................................................................................................................... iv	  Table of Contents ........................................................................................................................ vii	  List of Tables ................................................................................................................................ xi	  List of Figures .............................................................................................................................. xii	  List of Abbreviations ................................................................................................................. xiv	  Acknowledgements .................................................................................................................... xvi	  Dedication ................................................................................................................................. xviii	  Chapter 1: Introduction ................................................................................................................1	  1.1	   Cellular Ca2+ Signaling ................................................................................................... 1	  1.2	   Ryanodine Receptor Calcium Channels ......................................................................... 8	  1.3	   Cardiac Ryanodine Receptors ....................................................................................... 13	  1.4	   RYR2 and Excitation Contraction ................................................................................ 20	  1.4.1	   Hypothesis: RYR2 is required for cardiac function. ............................................. 24	  1.5	   RYR2 and Heart Rate ................................................................................................... 24	  1.5.1	   Hypothesis: RYR2 is required for cardiac heart rate and rhythm. ........................ 33	  1.6	   RYR2 and Cardiac Metabolism .................................................................................... 34	  1.6.1	   Hypothesis: RYR2 is required for mitochondrial Ca2+ uptake and oxidative metabolism ............................................................................................................................ 41	  1.7	   RYR2 and Cell Death ................................................................................................... 42	  1.7.1	   Hypothesis: RYR2 is required for cell survival. ................................................... 46	    viii 1.8	   RYR2 and Cardiac Hypertrophy ................................................................................... 47	  1.8.1	   Hypothesis: RYR2 is required for cardiac hypertrophy ....................................... 50	  1.9	   RYR2 and Heart Disease .............................................................................................. 50	  1.9.1	   Hypothesis: Decreases in Ryr2 expression cause dilated cardiomyopathy .......... 52	  Chapter 2: Materials and Methods ............................................................................................54	  2.1	   Experimental Animals .................................................................................................. 54	  2.2	   In Vivo Analysis of Cardiac Function and Heart Rate .................................................. 55	  2.3	   Ex Vivo Analysis of Cardiac Function and Metabolism ............................................... 56	  2.4	   Light and Electron Microscopy in the cRyr2KO Model ............................................... 56	  2.5	   Cardiomyocyte Ca2+ and Functional Measurements in the cRyr2Δ50 Model .............. 58	  2.6	   Gene and Protein Expression Analysis ......................................................................... 59	  2.7	   Metabolomics ................................................................................................................ 62	  2.8	   Transcriptomics ............................................................................................................. 63	  2.9	   Proteomics ..................................................................................................................... 64	  2.10	   Statistical Analysis ........................................................................................................ 65	  Chapter 3: Cardiac RYR2 Controls Heart Rate and Rhythmicity in Adult Mice ................66	  3.1	   Chapter Summary ......................................................................................................... 66	  3.2	   Introduction ................................................................................................................... 67	  3.3	   Results ........................................................................................................................... 69	  3.3.1	   Acute reductions in Ryr2 mRNA and protein in conditional knockout mice ....... 69	  3.3.2	   Sudden cardiac death in cRyr2KO mice ............................................................... 70	  3.3.3	   Heart function in cRyr2KO mice .......................................................................... 72	  3.3.4	   Loss of Ryr2 reduces heart rate and results in severe arrhythmias ....................... 74	    ix 3.4	   Discussion ..................................................................................................................... 78	  Chapter 4: Cardiomyocyte ATP Production, Metabolic Flexibility, and Survival Require Calcium Flux Through Cardiac Ryanodine Receptors In Vivo ..............................................83	  4.1	   Chapter Summary ......................................................................................................... 83	  4.2	   Introduction ................................................................................................................... 84	  4.3	   Results ........................................................................................................................... 86	  4.3.1	   Ryr2 deletion reduces Ca2+ flux in cardiac myocytes ........................................... 86	  4.3.2	   RYR2 supports ATP production in vivo ............................................................... 87	  4.3.3	   Ryr2 deletion leads to a hypoxic/ischemic state ................................................... 89	  4.3.4	   Reprogramming of metabolism in cRyr2KO hearts .............................................. 91	  4.3.5	   Ryr2 deletion induces programmed cell death ...................................................... 93	  4.3.6	   Loss of cardioprotective pathways in cRyr2KO hearts ......................................... 95	  4.3.7	   Ryr2 deletion causes cardiac hypertrophy and fibrosis ......................................... 95	  4.4	   Discussion ..................................................................................................................... 98	  Chapter 5: Cardiac RYR2 Specifically Promotes Cardiac Glucose Oxidation ...................104	  5.1	   Chapter Summary ....................................................................................................... 104	  5.2	   Introduction ................................................................................................................. 105	  5.3	   Results ......................................................................................................................... 107	  5.4	   Discussion ................................................................................................................... 122	  Chapter 6: Conclusion ...............................................................................................................128	  6.1	   Summary ..................................................................................................................... 128	  6.2	   General Caveats .......................................................................................................... 129	  6.3	   RYR2 and Excitation Contraction Coupling .............................................................. 132	    x 6.4	   RYR2 and Heart Rate ................................................................................................. 134	  6.5	   RYR2 and Metabolism ............................................................................................... 137	  6.6	   RYR2 and Cell Death ................................................................................................. 143	  6.7	   RYR2 and Hypertrophy .............................................................................................. 145	  6.8	   RYR2 and Cardiac Pathology ..................................................................................... 146	  6.9	   Overall Significance .................................................................................................... 150	  Bibliography ...............................................................................................................................153	     xi List of Tables Table 2.1. Sybr Green RT-qPCR probes………………………………………………………..60 Table 2.2. Commercially available Taqman RT-qPCR assays…………………………………61 Table 2.3. Custom Taqman RT-qPCR assays…………………………………………………..61 Table 2.4. Antibodies used in western blot experiments………………………………………..62   xii List of Figures  Figure 3.1. Acute, cardiac-specific Ryr2 gene ablation in mice…………………………………71 Figure 3.2. Conditional Ryr2 knockout is tissue specific and not compensated for by other SR Ca2+ release channels..…………………………………………………………………………...72 Figure 3.3. Conditional Ryr2 knockout mice rapidly lose cardiac function……………………..73 Figure 3.4. Conditional Ryr2 knockout hearts display reduced cardiac function………………..74 Figure 3.5. cRyr2 knockout hearts exhibit bradycardia and arrhythmias………………………..76 Figure 3.6. cRyr2 knockout mice exhibit bradycardia and arrhythmias in vivo…………………77 Figure 4.1. Acute, cardiac-specific Ryr2 gene ablation without compensation from related Ca2+ channels…………………………………………………………………………………………..88 Figure 4.2. Loss of RYR2-mediated Ca2+ flux reduces mitochondrial metabolism and steady-state ATP levels……………………………………………………………………………….....89 Figure 4.3. Acute Ryr2 ablation causes a hypoxia-like cellular state……………………………90 Figure 4.4. Conditional Ryr2 knockout results in metabolic reprogramming at the transcriptional level………………………………………………………………………………………………92 Figure 4.5. Acute Ryr2 ablation causes calpain-10 dependent programmed cell death ...............94 Figure 4.6. Cardiac hypertrophy and fibrosis in cRyr2KO hearts………………………………..97 Figure 5.1. Generation of an inducible, heart-specific Ryr2Δ50 knockout model with partial RYR2 ablation………………………………………………………………………………….108 Figure 5.2. Altered cytosolic and mitochondrial Ca2+ homeostasis in cRyr2Δ50 cardiomyocytes............................................................................................................................109 Figure 5.3. Cardiac function and heart rate in cRyr2Δ50 mice………………………………....111   xiii Figure 5.4. Metabolomic and working heart analyses of cRyr2Δ50 mice reveal a specific defect in glucose oxidation, associated with hyper-phosphorylation of PDH…………………………114 Figure 5.5: Proteomic analysis of cRyr2Δ50 hearts…………………………………………….116 Figure 5.6: Parallel analysis of functional protein categories in cRyr2Δ50 hearts……………..117 Figure 5.7: The Proteome of the electron transport chain, ATP synthase complex, and other metabolic effectors in cRyr2Δ50 hearts………………………………………………………...118 Figure 5.8: Transcriptomics analysis of cRyr2Δ50 hearts………………………………….…..121      xiv List of Abbreviations  ATP: Adenosine Triphosphate ATPase: Mitochondrial ATP synthase CaMK: Calmodulin dependent kinase CoA: Coenzyme A CPVT: Catecholaminergic polymorphic ventricular tachycardia DCM: Dilated cardiomyopathy ECC: Excitation-contraction coupling ECG: Echocardiography EM: Electron microscopy ER: Endoplasmic reticulum ETC: Electron transport chain FBP2: Fructose-bisphosphatase 2 FKBP: FK506 Binding Protein  HCN: Hyperolarzation-activating, inwardly rectifying cation channel HIF: Hypoxia inducible factor IP3: Inositol 1,4,5-triphospahate  IP3R: Inositol 1,4,5-triphospahate receptor MCU: Mitochondrial calcium uniporter MPTP: Mitochondrial permeability transition pore MLC: Myosin light chain NADH: Nicotinamide adenine dinucleotide   xv NCX: Na+/Ca2+ ion exchangers PKA: Protein kinase A PMCA: Plasma membrane calcium ATPase PP1: Protein phosphatase 1 PP2A: Protein phosphatase 2A PDE4D3: Phosphodiesterase 4D3 PDH: Pyruvate dehydrogenase PDK: Pyruvate dehydrogenase kinase PDP: Pyruvate dehydrogenase phosphatase PFK1: Phosphofructokinase 1 RYR: Ryanodine receptor  RYR2: Ryanodine receptor 2 SCN5: Voltage-gated sodium channel Scn5 SERCA: Sarco/endoplasmic reticulum ATPase SR: Sarcoplasmic reticulum TCA: Tricarboxylic acid cycle, aka the ‘citric acid cycle’ or ‘Krebs cycle’ VDAC: Voltage dependent anion carrier VGCC: Voltage-gated calcium channel       xvi Acknowledgements  I owe many thanks to Heather Bround for her love and support, as well as generally putting up with my unorthodox career choice. Novelists always thank their partners as if being in a relationship with a writer is The Worst Thing, and having actually written something of substantial length I can see why. Sorry for the writing absence and thank you for understanding. I love you and couldn’t ask for a better partner. Special thanks are owed to my parents for their love and their support of my education. You made education a priority for me and made attending university attainable. I could not have gotten this far without you. I also owe buckets of gratitude to my PhD supervisor Professor James D Johnson for his mentorship and giving me the opportunity to work on this project. He has been generous with his time, resources, and knowledge. I especially appreciate that he was willing to take a chance on this foray into cardiac biology. I am leaving his lab a much better scientist than when I entered and have really enjoyed this time working with him. I would like to thank Professor Michael A Allard, Professor Edwin D Moore, and Professor Dan S Luciani, for being members of my PhD committee and for their time, guidance, and expertise. In addition, I thank Professor Roger W Brownsey for his time and his profound knowledge on the minutiae of biochemical processes. It has been a pleasure. I would like to thank my various collaborators: Dr. Rich Wambolt, Dr. Parisa Asghari, Dr. Razvu Albu, Dr. Don McAfee, Dr. Marc Pourrier, Dr. Nichollas Scott, Dr. Jun Han, Lubos Bohunek, Marjorie Philit, Dr. Jerzy Kulpa, Dr. Brian Rodrigues, Dr. David Fedida, Dr. Christoph Borchers, Dr. Leonard Foster, Dr. Thibault Mayor, and Dr. Tim Kieffer. Thank you all for your   xvii contributions to the work presented in this thesis, your shared expertise and instruction, and your generosity. I could not have done this research without you. I would like to thank all of the members of the Johnson Lab during my time there. The quality of my graduate school experience is in large part to the relationships and scientific interaction I ave had with this group. I have enjoyed being part of this team. I would particularly like to thank Dr. Carol Yang, Dr. Tobias Albrecht, Dr. Nicole Templeman, Sarah Gage, and ‘honorary’ lab members Dr. Blair Gage and Anna D’Souza for their friendship and scientific interaction. You made this whole process fun. I would also like to thank Dr. Scott Covey for his undergraduate mentorship and his help reaching graduate school and Dr. Jason Reed for access to teaching opportunities. I would like to thank Dr. Navkiran Gill, Dr. Emily Mace, Dr. Fumio Takei, and Dr. B Brett Finlay for the undergraduate research opportunities that convinced me to pursue research as a career.  I would also like to thank Andrew Bround, Kathleen Fraser, and Alexandra Capistrano for their friendship and support.    xviii Dedication   For Shiloh.  1 Chapter 1:  Introduction  1.1   Cellular Ca2+ Signaling One of the key concepts in biology is ‘homeostasis’: the idea that living organisms regulate their internal environment in an optimal set of conditions for biological processes (Cannon, 1926). Cells must have mechanisms to control their internal environment and ensure the chemical and thermodynamic requirements of their biological processes are constantly met. Since biological organisms exist in a non-static environment that can change in potentially disruptive ways, cells must have the capacity to adapt to changes in their environment to maintain homeostasis. Therefore, cells require a means to sense their environment, communicate this information into the cell, and activate a variety of adaptive cellular responses. This process of homeostasis becomes more complex in multi-cellular organisms. In multi-cellular organisms, many cells work in concert to maintain whole organism homeostasis. Many complex organisms also display cellular specialization, where discrete groups of cells perform specific functions. In this situation, it becomes critical for cells to communicate with one another to coordinate collective cellular responses and to regulate specialized functions. This involves many of the same principles of environment sensing, internal communication, and cell response as general homeostasis. The mechanism by which cells activate cellular processes in response to extracellular cues is called ‘cellular signaling’ (Johnson et al., 2014). While there is a vast diversity of processes that fall under this label, the majority of signaling involves three general components: a sensor protein which recognizes a cue and initiates a cellular signal, a second messenger which disseminates the signal to its intracellular target effector, and effector proteins that are activated   2 by the second messenger and initiate a cellular response. With this basic system, cells are able to directly couple their behavior to their external environment or to respond to signals originating elsewhere within a multicellular organism.  Sensor proteins or protein complexes are as diverse as the cues that activate them. There are signaling sensors which recognize changes in temperature (Peier et al., 2002), respond to protein hormones (Hubbard, 2013), detect changes in voltage across cellular membranes (Catterall, 2000), sense metabolic state (Tucker et al., 1998), measure mechanical forces (Kwan et al., 2006), detect photons of light (Khorana, 1992), and respond to a vast array of other conditions or molecules. Despite this complexity, there are some common features: sensor proteins or complexes will contain a region that binds a target ligand or interacts with an environmental condition, and a region that is able to stimulate the production or release of a second messenger molecule which will go onto activate a cellular response (Johnson et al., 2014).  Second messengers are the media of cellular signaling: they are the physical substance conveying the message of a signal to its target responders. Generally speaking, they are small molecules that can rapidly diffuse in the cytoplasm to propagate a signal (Berridge et al., 2000; Johnson et al., 2014). These small molecules are either rapidly synthesized upon activation of a signaling cascade (Berridge, 1993; Eichmann and Lass, 2015; Rosenbaum et al., 2009) or admitted into the cellular milieu from either the extracellular space or an intracellular storage location (Berridge et al., 2000; Johnson et al., 2014). Once released into the cytosol, second messengers will rapidly diffuse and bind to their target receptors, activating them and stimulating a cellular response. Second messenger diffusion is restricted by buffering molecules which bind the signaling molecules (Berridge et al., 2000; Gilabert, 2012; Mika et al., 2012), and by terminating enzymes which breakdown or remove second messengers from the milieu, limiting   3 their spread and helping to end the cellular signal (Berridge et al., 2000; Mika et al., 2012). Cellular signaling is therefore shaped by the competing forces generating, restricting, and removing a second messenger during a signaling event. A variety of molecules can act as second messengers, each with different kinetics, physical properties, and signaling partners.  Signaling effectors are the cellular machinery that generate a biological response to a signal. Again, there is an impressive diversity of effectors, with different activating second messengers, a variety of biochemical functions, and a wide range of different responses. There are some common themes. Many signaling effectors are protein kinases or phosphatases which modify the phosphorylation state of downstream proteins in response to a signaling event, altering their behavior or activity (Hunter, 1995). Many other signaling effectors are involved in transcription factor activation or localization, which functions to alter the gene expression patterns within a cell (Spitz and Furlong, 2012). Collectively then, cellular signaling effectors provide cells a way to modulate protein function and transcription to extracellular cues and signals. One of the most conserved cellular signaling systems uses calcium as a second messenger (Berridge et al., 2000; Johnson et al., 2014). In biological systems, calcium exists as a cation carrying a positive two charge (Ca2+). Perhaps due to the fact that Ca2+ precipitates phosphate ions, a circumstance that would be disadvantageous for cells using high-energy phosphates as metabolic currency, Ca2+ has largely been excluded from the basal cellular milieu (Berridge et al., 2000). As a result cytosolic Ca2+ concentration is aggressively maintained at a very low level, ~100 nM (Berridge et al., 2000; Pozzan et al., 1994), which creates a driving force for Ca2+ entry into the cytosol and allows the cation to rapidly diffuse through the cytoplasm (Berridge, 1997). The low basal concentration of cytosolic Ca2+ is also significant as it allows for a wide working range where target effectors with significantly different sensitivities to Ca2+ can grade cellular   4 responses through amplitude encoding (Berridge et al., 2000; Dolmetsch et al., 1997; Goll et al., 2003). The specific localization of Ca2+ sources and effectors, and the discrete kinetics of Ca2+ release and removal, also grant Ca2+ signaling a degree of spatial (Berlin et al., 1994; Gilabert, 2012; Matthews et al., 1994; Pozzan et al., 1994; Thomas et al., 1996) and frequency encoding (Dolmetsch et al., 1997; 1998) which allow for more fine-tuned and specific signaling. This makes Ca2+ a robust second messenger capable of both large global cellular responses and localized, discrete signaling events. Ca2+ signaling is involved in many cellular responses in a variety of cell types and tissues. As a result, it is a complex process with many context specific nuances. However, Ca2+ signaling generally falls into two signaling paradigms: an extracellular Ca2+ import or an intracellular Ca2+ release. In the case of extracellular Ca2+ signaling, the cell takes advantage of the extreme Ca2+ gradient between the extracellular space (which in mammals is rigorously maintained to be between 2.2 and 2.7 mM (Brown, 1991)) and the cytosol to passively allow Ca2+ to rush into the cell in response to a stimulus. In this case, the cell simply opens a Ca2+ channel located in the plasma membrane to give the ion a pathway through the impermeable lipid bilayer and into the cell (Berridge et al., 2000). This can occur in response to an extracellular ligand or condition directly in the case of the transient receptor potential (TRP) channel family which open for a wide variety of stimuli and directly facilitate Ca2+ uptake (Minke and Cook, 2002; Nieto-Posadas et al., 2011; Ramsey et al., 2006). Ca2+ entry can also occur in response to voltage changes across the cellular membrane via voltage-gated Ca2+ channels (VGCCs) (Catterall, 2011). In this circumstance, a charge differential across the plasma membrane interacts with voltage sensing domains in VGCCs to allosterically cause the channel to open and permit Ca2+ entry into the cell (Catterall, 2000). VGCCs come in three main classes: CaV1 (or L-type) Ca2+ channels which   5 open at high voltage differentials and permit relatively large, sustained Ca2+ signals; CaV3 (or T-type) channels which open at lower voltage differences, but which only permit short, small Ca2+ entry events; and CaV2 channels which have a voltage and Ca2+ conductance intermediate to the other two (Catterall, 2011). Regardless of the activating stimuli, Ca2+ entry is eventually terminated by the closing of the entry channel and the export of Ca2+ from the cytosol back to the extracellular space via Na+/Ca2+ ion exchangers (NCX), which use the cellular Na+ gradient to rapidly remove cytosolic Ca2+ (Brini and Carafoli, 2011; Philipson and Nicoll, 2000; Reeves and Condrescu, 2008), and the plasma membrane calcium ATPase (PMCA), which hydrolyzes ATP to pump Ca2+ out of the cell (Brini and Carafoli, 2011; Di Leva et al., 2008; Giacomello et al., 2013).  In the case of intracellular Ca2+ release, Ca2+ is mobilized from internal stores to initiate a cellular response. Cells sequester Ca2+ in intracellular organelles, most importantly the sarcoplasmic/endoplasmic reticulum (SR/ER) where Ca2+ plays a role in protein folding and cellular signaling (Berridge et al., 2000). Cells maintain SR/ER Ca2+ at ~100 µM, which means that Ca2+ stored in this organelle also has a steep concentration gradient driving it’s entry into the cytosol (Pozzan et al., 1994; Stutzmann and Mattson, 2011). Similar to extracellular Ca2+ entry, SR/ER Ca2+ release is governed by the opening of protein channels which allow Ca2+ to cross the ER/SR membrane and passively diffuse into the cytoplasm (Berridge et al., 2000; Pozzan et al., 1994). In the case of SR/ER there are two main types of Ca2+ release channels: the ryanodine receptors (RYR) channel family and the Inositol 1,4,5-triphospahate receptor (IP3R) channel family. The RYR family are Ca2+ induced Ca2+ release channels that are activated mainly by binding cytosolic Ca2+ and which release a large amount of ER/SR Ca2+ into the cytosol when activated (Lanner et al., 2010; Van Petegem, 2015). RYR are often functionally coupled to   6 voltage gated Ca2+ channels and serve as signal amplifiers which convert small, localized Ca2+ signals into large global ones (Lanner et al., 2010). IP3R channels release ER/SR Ca2+ in response to binding the lipid messenger inositol 1,4,5-triphosphate, which is generated by phospholipase C in response to extracellular cues (Berridge, 2009; Foskett et al., 2007). Intracellular Ca2+ stores are reloaded by the sarco/endoplasmic reticulum ATPase (SERCA) family of Ca2+ pumps (Brini et al., 2012; Møller et al., 2010). These proteins consume ATP to rapidly pump Ca2+ ions back into the ER/SR which reloads the Ca2+ store and contributes to terminating Ca2+ signals (Møller et al., 2010; Pozzan et al., 1994). Therefore, the intracellular Ca2+ release of the SR/ER in many respects mirrors the extracellular Ca2+ signaling of the plasma membrane, but in a different location. While other organelles can participate in Ca2+ signaling, notably endosomes via two pore channels (Calcraft et al., 2009), the ER/SR signaling system is in most situations the largest contributor to intracellular Ca2+ signaling. Ca2+ signaling is involved in a wide variety of processes and, as such, there are many target effectors with unique chemical and signaling properties. However, there are some common structural motifs and major proteins worth consideration. The vast majority of Ca2+ sensitive proteins interact with the cation via the ‘EF-hand’ protein domain (Grabarek, 2006; Kawasaki et al., 1998). EF-hands are modular Ca2+ binding domains that contain an ion binding pocket between two alpha helixes, oriented in such a way that they resemble ‘hands’ (hence the name) (Grabarek, 2006; Kawasaki et al., 1998). When Ca2+ binds in this domain, it causes a change in the conformation of the EF-hand, which in turn causes an allosteric change in its host protein, frequently uncovering an active site or binding domain that changes the activation state of the host protein (Grabarek, 2006; Kawasaki et al., 1998). While many proteins contain their own EF-hand domains (Grabarek, 2006), many other Ca2+ sensitive proteins take advantage of a Ca2+   7 sensor protein known as calmodulin (Finn and Forsén, 1995). Calmodulin is a small, EF-hand domain containing protein that changes from a linear, inactive conformation to a globular conformation with exposed protein binding domains when exposed to Ca2+ (Finn and Forsén, 1995). Many proteins, rather than directly sensing Ca2+, rely on calmodulin-binding for activation or inactivation, and are therefore indirectly sensitive to Ca2+ (Hoeflich and Ikura, 2002; Tidow and Nissen, 2013). Two of the most common and important Ca2+ dependent effector classes are the calmodulin dependent kinase (CaMK) family (Wayman et al., 2011), which phosphorylates target proteins in response to a Ca2+ signal, and the calmodulin dependent phosphatase, calcineurin family (Rusnak and Mertz, 2000), which removes regulatory phosphate groups from proteins in a Ca2+ dependent manner. These two proteins are able to regulate a host of downstream targets, altering the activity of enzymes, changing the functional parameters of other proteins, and altering the localization and activity of a variety of transcription factors. While specific signaling targets differ, many Ca2+ dependent cellular responses involve these proteins or domains, or the overall chemical trends of these examples. Ca2+ is a virtually ubiquitous second messenger throughout biology and thousands of proteins and enzymes are sensitive to Ca2+, either directly or indirectly. As such, Ca2+ signaling controls a wide variety of cellular responses in many different cell types in mammalian biology. Ca2+ signaling can alter universal cellular processes such as gene expression (Dolmetsch, 2003; Heineke and Molkentin, 2006), protein regulation (Rusnak and Mertz, 2000; Wayman et al., 2011), metabolism (Denton and McCormack, 1990), cell differentiation and proliferation (Capiod, 2013; Oh-hora, 2009), and programmed cell death (Luciani et al., 2008). Ca2+ signaling is also central to specialized cellular processes such as hormone exocytosis (Kasai et al., 2010; Petersen, 1992), myocyte cell contraction (Bers, 2001), neurotransmitter releases (Neher and   8 Sakaba, 2008), immune cell adhesion and motility (Newton and Dixit, 2012), setting heart rate and rhythmicity (Monfredi et al., 2013), and a host of other critical cellular processes. Ca2+ signaling is a fundamental aspect of biology and a critical part of cellular homeostasis and the coordinated function of multicellular organisms. Ca2+ signaling, therefore, is a critical process to study. Given it’s central role in regulating a variety of cellular processes and responses, understanding the role of Ca2+ signaling in any given system is an important part of fundamentally understanding biology. Moreover, since Ca2+ signaling and homeostasis is often disrupted in various disease states, such as diabetes (Arruda and Hotamisligil, 2015), heart disease (Gorski et al., 2015), and Alzheimer’s disease (Demuro et al., 2010; Woods and Padmanabhan, 2012), it becomes important to understand how Ca2+ signaling contributes to pathophysiology. Doing so will not only uncover valuable insight into disease, but may also provide novel therapeutic targets, such as has been found VGCC channel blockers which are used to treat hypertension, epilepsy, and chronic pain (Zamponi et al., 2015). My specific goal in this thesis is to better understand Ca2+ signaling in the cardiomyocyte and to study how the professional, contractile Ca2+ signaling of the cell also regulates a variety of other processes critical for cardiac biology and health.  1.2   Ryanodine Receptor Calcium Channels The history of the ryanodine receptor Ca2+ channel is intimately connected to the plant toxin ryanodine. Ryanodine is a plant alkaloid molecule found in the stem and roots of the plant Ryania speciosa vahl, a species native to Trinidad (Jenden and Fairhurst, 1969). Apparently, there was a period when crude Ryania extracts were used commercially as an insecticide due to the compounds extensive toxicity to insects (Heal, 1949). In the 1920s, experiments were   9 conducted which explored the toxicity of crude Ryania extracts in vertebrates, and noted that the mixtures caused respiratory paralysis and death in frogs, mice, cats, dogs, but not fish (Procita, 1958). In 1948 pure ryanodine was isolated from Ryania extracts, crystallized and chemically characterized (Rogers et al., 1948). It was subsequently found, in the 1950s, that ryanodine was the key toxic component of Ryania extracts and that administering ryanodine to mammals at extremely low doses caused respiratory paralysis and collapse of cardiac function (Procita, 1958) as well as irreversible contracture in isolated muscle fibers from frogs and mammals (Blum et al., 1957).  What followed was a wave of science exploring the toxicity of ryanodine and attempting to discover the mechanism of the molecules action on skeletal and cardiac muscles (Jenden and Fairhurst, 1969). In hindsight, it is an interesting body of literature, since it was confounded by differences in model organisms and the dual action of ryanodine, which acts as an agonist of the Ca2+ channel in nanomolar concentrations and an inhibitor in micromolar doses (Sutko et al., 1997). Eventually, in the 1980s it was realized that ryanodine had dual effects on SR Ca2+ release, and the compound was binding some sort of unknown channel or protein in the SR of striated muscle tissues associated with ‘junctional foot’ structures observed in electronmicrographs (Chamberlain et al., 1984b; 1984a; Fleischer et al., 1985; Meissner, 1986). Leveraging this knowledge, work done mainly by the groups of Fleisher and Meissner was able to isolate the so called “ryanodine receptor” from cardiac and skeletal muscle SR (Anderson et al., 1989; Inui et al., 1987a; 1987b; Lai et al., 1988) and to demonstrate this was indeed the Ca2+-induced Ca2+ release channel involved in muscle contraction. Which is how a plant based insecticide was used, over the course of decades of scientific work, to discover one of the most important proteins in myocyte biology.   10 Subsequent decades of work have revealed that there are three isoforms of ryanodine receptor in the mammalian ryanodine receptor family: RYR1, RYR2, and RYR3 (Lanner et al., 2010). All three isoforms are massive protein channels that function as ER/SR Ca2+ release channels which are activated by binding cytosolic free Ca2+ (Lanner et al., 2010). Since RYRs release relatively large amounts of Ca2+ (~100 pS Ca2+) in response to Ca2+ (Ondrias et al., 1996), they function as signal amplifiers, increasing the amplitude of smaller Ca2+ signals. In addition, since individual RYR channels are able to activate one another, either via Ca2+ release (Berridge, 1997) or allosteric interactions (Marx et al., 2001a; Porta et al., 2012), RYRs are able to propagate Ca2+ signals within a cell and are able to turn local Ca2+ signaling events into global ones (Berridge, 1997; Cheng et al., 1996). RYRs can also function as docking sites for numerous regulatory and structural molecules (Van Petegem, 2015), allowing the channels to integrate and respond to a variety of cellular signals and to potentially function as structural nucleation points. RYRs are therefore complex signaling proteins poised to contribute to a wide variety of Ca2+ dependent processes. RYR channels are found in a wide variety of cell types including immune cells (Hosoi et al., 2001), endocrine cells (Luciani et al., 2008; Santulli et al., 2015), neurons (Lai et al., 1992), and skeletal and cardiac myocytes (Bers, 2002; Inui et al., 1987a; Yang et al., 2001). Type 1 ryanodine receptors (RYR1) are generally considered the ‘skeletal muscle isoform’, since it is the primary isoform found in this tissue (Endo, 2009; Inui et al., 1987b). The type two ryanodine receptor (RYR2) isoform is often referred to as the ‘cardiac isoform’ since it is the dominant member of the protein channel in cardiac myocytes (Inui et al., 1987a; Lehnart et al., 2004a). This is somewhat misleading as both RYR1 and RYR2 are found in a wide variety of other cell types. Notably, RYR2 is found in neurons where it is widely expressed at substantial levels   11 (Hidalgo et al., 2015; Kurokawa et al., 2011; Lai et al., 1992). RYR3 also has a wide tissue distribution and is most highly expressed in the brain (Sorrentino, 2003). Collectively the RYRs are a common feature of most mammalian cell types and can be highly expressed and centrally involved in major cellular processes or low level, modest contributors to cellular function and housekeeping processes. Structurally, RYRs are ER/SR spanning ion channels. While there are some key structural differences between isoforms, overall RYRs share several common structural elements. RYRs are 2.2 MDa proteins comprised of four identical monomers which each contain two β-trefoil domains, one armadillo repeat domain, three SpIA kinase ryanodine receptor (SPRY) domains, a Ca2+ sensing EF-hand domain, as well as disordered isoform divergent regions, general structural regions specific to RYRs, and 6 transmembrane helices that collectively form the channel pore (Efremov et al., 2015; Van Petegem, 2015; Yan et al., 2015; Zalk et al., 2015). When assembled, the RYR tetramer has a symmetrical ‘mushroom’ shape, with a relatively small ER luminal region, a transmembrane region, and a large cytosolic ‘cap’ that makes up the majority of the protein and provides surface area for protein interactions (Serysheva et al., 2008; Van Petegem, 2015). The cytosolic ‘cap’, is further divided by structural biochemists into three regions called the ‘clamps’, ‘handle’ and ‘central rim’ (Serysheva et al., 2008; Van Petegem, 2015). The exact molecular identity of the RYR cytosolic Ca2+ sensing domain remains controversial. Some evidence suggests that the EF-hand domain, located in the central domain of the protein and physically located near the C-terminal domain of each monomer, may act as a Ca2+ sensor (Efremov et al., 2015; Xiong et al., 2006), although other work disputes this, showing that deletion of the EF-hand regions in RYR1 and RYR2 does not affect cytosolic Ca2+ activation but instead may be involved in luminal Ca2+ release (Fessenden et al., 2004; Guo et al., 2015). Other   12 work has suggested that a nearby glutamate residue may also bind Ca2+ and participate in cytosolic channel activation (Li and Chen, 2001). RYRs also posses the ability to sense and react to changing SR/ER luminal Ca2+ levels, and data suggests that both the RYR2 EF-hand domains as well as a glutamate residue in the pore-forming region are involved in luminal Ca2+ sensing (Chen et al., 2014; Guo et al., 2015). RYR structure is an area of very active research, so the current understanding of the relationship between channel structure and function is rapidly improving. Another important RYR structural consideration is that individual RYR channels are able to physically interact with one another to form lattices via the ‘clamp’ region of their cytosolic portion (Franzini-Armstrong et al., 1999; Marx et al., 2001a; 1998; Porta et al., 2012; Yin and Lai, 2000; Yin et al., 2005b). Under certain conditions, RYRs will spontaneously form a ‘checkerboard’ lattice, and it is believed that this structural arrangement potentiates collective gating, where groups of RYRs can simultaneously signal via allosteric interaction (Franzini-Armstrong et al., 1999; Marx et al., 1998; 2001a; Porta et al., 2012; Yin and Lai, 2000; Yin et al., 2005b). It has also been shown that, at least for RYR2 in cardiomyocytes, that this arrangement is context dependent and dynamic and that RYR2 will switch between a cooperative checkerboard and a less interactive, denser conformation in a Mg2+ and/or phosphorylation dependent mechanism (Asghari et al., 2014; Yin et al., 2005a). In this sense RYRs function both as individual channels and structurally as larger channel complexes with the potential for collective signaling. RYRs experience large-scale motions during their activation that cause dramatic conformational changes throughout the channels structure (Van Petegem, 2015). Recent work has suggested that RYR channels are held closed by an interaction between the EF-hand domain,   13 the flexible C-terminal domain, and a voltage sensor like domain, located in a loop between transmembrane domains (Efremov et al., 2015; Yan et al., 2015; Zalk et al., 2015). It is believed that Ca2+ binding to the cytosolic Ca2+ sensor, whatever its molecular identity, disrupts this complex which causes the channel to open (Efremov et al., 2015; Yan et al., 2015; Zalk et al., 2015). The channel can also open, or sensitized to open, by the luminal Ca2+ sensor, which involves residues at the bundle crossing on the ER/SR side of the channel pore and possibly the EF-hand domains (Chen et al., 2014; Guo et al., 2015; Qin et al., 2008). Regardless of the site of activation, it is thought that channel activation induces large-scale conformational changes that result in the channel pore swelling and creating a path for Ca2+ to flood into the cytosol. This motion also causes the structures immediately around the pore to move apart and the cytosolic ‘cap’ to ‘tilt’, which moves the ‘central rim’ outward and causes the ‘clamps’ to bend towards the ER/SR (Van Petegem, 2015). Since the ‘clamp’ regions are the points where individual RYR channels are thought to interact with one another, it has been proposed that the allosteric motion in this region is the mechanism by which the activation state of one channel is conveyed to channels within the same array (Van Petegem, 2015). Thus, RYRs utilize conformational changes to drive alterations within single channels and potentially to communicate activation states within coordinated groups of channels.  1.3   Cardiac Ryanodine Receptors Ryanodine Receptor 2 (RYR2) is the primary isoform of RYR in the heart (Lehnart et al., 2004a). While this protein has a wide tissue distribution (Lanner et al., 2010), it was originally discovered in the heart (Inui et al., 1987a), and is expressed at a much higher level here than in any other tissue. RYR2 is activated primarily by cytosolic free Ca2+, which in the heart, is   14 provided primarily by the cardiac L-type voltage gated Ca2+ channel (Bers, 2001; Lehnart et al., 2004a). RYR2 shows a biphasic sensitivity to Ca2+, where cytosolic Ca2+ concentrations of ~1 µM are required to activate the channel and cytosolic concentrations greater than ~1 mM prevent the channel from opening (Copello et al., 1997; Laver et al., 1995; Meissner et al., 1986). It is not commonly believed that electrochemical coupling plays a significant role in RYR2 activation (Bers and Stiffel, 1993; García et al., 1994; Perez et al., 2003). RYR2 is also sensitive to SR luminal Ca2+ concentration such that the channel has increased open probability at higher luminal Ca2+ concentrations and that RYR2 Ca2+ release can be triggered outright by so called ‘store overload’ when SR Ca2+ exceeds a critical threshold. (Chen et al., 2014; Guo et al., 2015; Qin et al., 2008). RYR2 is a very high conductance Ca2+ channel, with a single channel conductance of ~100-150 pS for Ca2+ (Tinker and Williams, 1992). This allows cardiac RYR2 to increase cytosolic Ca2+ concentration from ~100 nM to ~10 µM during peak Ca2+ signalling in cardiomyocytes (Bers, 2001). RYR2 is generally specific for Ca2+ ions, but may also transport Mg2+ ions under certain conditions (Tinker and Williams, 1992). The exact mechanism governing channel closing remains controversial, but is thought to involve either inhibition from local increases in cytosolic Ca2+ levels caused by the RYR2 channels themselves (Lukyanenko et al., 1998; Sham et al., 1998) or by depletion of SR Ca2+ levels by channel activity (Bers and Shannon, 2013; Györke and Györke, 1998; Sitsapesan and Williams, 1994). However this latter theory is unlikely, since RYR2 channels lacking the putative SR Ca2+ sensor do not demonstrate a loss of cytosolic Ca2+ activation or RYR2 refractoriness (Guo et al., 2015). RYR2 channels represent the main SR Ca2+ release channel of the cardiomyocyte and play a major role in cardiac Ca2+ signaling.   15 Structurally RYR2 shares many of the features common to all members of the RYR protein family (Van Petegem, 2015). RYR2 is a 2.2 MDa, homotetramer, with the common RYR protein ‘mushroom’ shape and four-fold symmetry (Van Petegem, 2015). To date the best high resolution EM studies have only been conducted on RYR1 (Efremov et al., 2015; Yan et al., 2015; Zalk et al., 2015), the skeletal muscle isoform, however RYR2 is predicted to share many of the same features by homology: the majority of the cytosolic cap, the probable cytosolic Ca2+ sensor, the six transmembrane domain channel pore, and the SR luminal Ca2+ sensor are all found in regions of very high sequence homology between isoforms (Van Petegem, 2015). The main structural difference between RYR2 and RYR1 is found in the D1, D2, and D3 divergent regions which has been implicated as the source of electromechanical coupling in RYR1 in skeletal muscle (Perez et al., 2003; Block et al., 1988), which is important for the sustained Ca2+ signals underlying the summation of tetanus in skeletal muscle contraction (Endo, 2009).  RYR2 possesses a large number of phosphorylatable sites, perhaps as many as 140 per channel tetramer (Van Petegem, 2015). While the vast majority of these sites have an undefined role in RYR2 function, it has been suggested that serine 2808 and/or serine 2030 are phosphorylated by PKA (Xiao et al., 2006; Zalk et al., 2007), the downstream kinase of beta-adrenergic signaling which increases channel opening probability, and that serine 2808 and serine 2814 are targeted by CaMKII for phosphorylation (Wehrens et al., 2004). RYR2 additionally has several residues that can be oxidized, nitrosylated, and glutathionylated, modifications that have been associated with altered channel gating and link cellular redox state to RYR2 function (Boraso and Williams, 1994; Donoso et al., 2011; Oda et al., 2015; Sánchez et al., 2005; Stoyanovsky et al., 1997; Xu et al., 1998). It has also been shown that RYR2 readily forms cooperative multi-channel lattice structures via ‘clamp’ region interactions, but it has been shown that these signaling units are   16 dynamic and are regulated by Mg2+ concentration and RYR2 regulatory phosphorylation state (Asghari et al., 2014; Marx et al., 2001a). Structurally, RYR2 is mostly understood via its regions of common homology with other RYRs, and further work is needed to understand what specific functional roles are linked to the divergent regions of the channel. RYR2 also interacts with a diverse array of regulatory proteins that modify the channel behavior of the RYR2 complex. The main region available for protein interaction is the cytosolic ‘cap’, which is not a single, solid object but rather a porous ‘vestibule’ that provides a large surface area for protein associations and interactions to occur (Bers, 2004; Van Petegem, 2015). These interactions would include the proteins involved in the regulatory phosphorylation sites discussed above: both PKA and CaMKII will co-immunoprecipitate with RYR2 as well as PP1, PP2A, and PDE4D3, the phosphatases which inactivate kinase signaling (Marx et al., 2001b; Ullrich et al., 2012; Wehrens et al., 2004; Zalk et al., 2007). Another regulatory interaction is with the ubiquitous Ca2+ sensing molecule calmodulin. One calmodulin protein is thought to bind per RYR2 subunit in both its Ca2+ bound and unbound state (Balshaw et al., 2001; Bers, 2004; Fuentes et al., 1994; Huang et al., 2012; Maier and Bers, 2002). Binding Ca2+-bound calmodulin has been demonstrated to decrease RYR2 open probability and increase the concentration of activating Ca2+ needed to activate the channel, potentially by restricting the motion of the ‘clamp’ regions of the channel ‘cap’ (Balshaw et al., 2001; Fuentes et al., 1994; Huang et al., 2012; Van Petegem, 2015). RYR2 also binds FK-binding proteins: both FKBP12 and FKBP12.6 are expressed in cardiac tissue and are able to bind the cytosolic ‘cap’ (Sharma et al., 2006; Van Petegem, 2015). Binding FKBP12 and FKBP12.6 has been shown to decrease RYR2 channel open probabilities, but also decrease the time spent in subconductance states, which combines to make RYR2 function with fewer, larger transients (Brillantes et al., 1994). It has also been   17 proposed that FKBPs may also be involved in the mechanism of coupled channel gating between RYR2 channels (Marx et al., 2001a). There is also evidence that RYR2 interacts with sorcin, a cytosolic EF-hand containing Ca2+ sensor protein (Farrell et al., 2003; Lokuta et al., 1997; Meyers et al., 1995). While the exact binding site remains unknown, there is evidence that sorcin reduces RYR2 open probabilities when cytosolic Ca2+ is greatly elevated (50-200 µM), and that this effect can be reduced by PKA signaling (Farrell et al., 2003; Lokuta et al., 1997; Meyers et al., 1995). RYR2 has also been shown to be affected by homer1c, an adapter protein that associates with a variety of ion channels, which has been shown to reduce RYR2 opening, dramatically shorten channel open times, and perhaps sensitize RYR2 to lower cytosolic Ca2+ levels (Pouliquin and Dulhunty, 2009; Pouliquin et al., 2009; Westhoff et al., 2003). It is currently unknown what regulates RYR2/homer1c interactions (Pouliquin and Dulhunty, 2009). RYR2 gating probability is also modified by binding cytosolic ATP, which increases channel open probability, and Mg2+ which reduces channel opening, either by competing for Ca2+ binding sites and/or decreasing cooperative activation within groups of RYR2 (Asghari et al., 2014; Meissner et al., 1986). Collectively these regulatory proteins alter RYR2 gating allowing the channel’s function to be tuned by a variety of other cellular signaling inputs providing the cell with an additional level of control and making RYR2 a major node of cellular signal integration. RYR2 also interacts with several proteins in the SR lumen that are important for RYR2 function and modulating channel behavior. In the SR lumen RYR2 forms a complex with the proteins junctin, triadin, and calsequestrin (Scriven et al., 2013). Cardiac calsequestrin 2 is a high capacitance (20 Ca2+ ions per protein), low affinity Ca2+ buffer (Km ~500 µM) that may provide a localized pool of rapidly mobilizable Ca2+ but may also play a role in stabilizing RYR2 function and channel refractoriness (Beard et al., 2004; Gaburjakova et al., 2012; Györke et al., 2004;   18 Mitchell et al., 1988). Junctin and triadin are thought to play a structural role in tethering calsequestrin molecules to RYR2 (Gaburjakova et al., 2012; Györke et al., 2004). Histidine rich Ca2+-binding (HRC) protein is another high capacitance, low affinity Ca2+ buffer that has been shown to bind triadin in the heart (Gregory et al., 2006; Lee et al., 2001). It is thought that CASQ2 and HRC combine to substantially increase the Ca2+ carrying capacity of cardiac SR to facilitate Ca2+ signaling (Bers, 2004; Scriven et al., 2013). However, it has recently been proposed that HRC, despite its structural similarities to CASQ2, may have a divergent role in regulating RYR2 gating, possibly promoting RYR2 Ca2+ release and reducing refractoriness (Liu et al., 2015). In this model CASQ2 and HRC may compete for degenerate binding sites on triadin in a frequency or Ca2+ concentration dependent manner where they have opposing effects on RYR2 Ca2+ release (Liu et al., 2015). This, then, may represent an additional mechanism for luminal Ca2+ regulation of RYR2 gating (Liu et al., 2015). Regardless, RYR2 has additional mechanisms to sense and respond to SR Ca2+ levels which means that it can integrate both cytosolic and SR luminal conditions to modulate its signaling behavior. RYR2 may also play a role as a structural molecule. Given the channel’s size and ability to interact with both cytosolic and SR luminal proteins, RYR2 is poised to function as a nucleation point for a variety of important cardiac structures (Bers, 2004; Scriven et al., 2013). Given that individual RYR2 bind one another to form signaling arrays (Asghari et al., 2014; Marx et al., 2001a), RYR2 clearly has an important role in maintaining Ca2+ signaling structures. In addition, RYR2 complexes are thought to interact with junctophilin, a protein that is involved in tethering the plasma membrane to the SR at signaling junctures (Jayasinghe et al., 2012; Scriven et al., 2013; Takeshima et al., 2000). While the exact relationship between RYR2 and cardiac junctophilin 2 remains unclear, in skeletal muscle RYR1, the skeletal muscle L-type VGCC, and   19 junctophilin 1 have been shown to physically interact to maintain the close association between these proteins (Golini et al., 2011). Knockdown of junctophilin 2 does significantly reduces co-localization between RYR2 and the cardiac L-type VGCC suggesting a similar role (van Oort et al., 2011). In addition there is some evidence that RYR2 may weakly interact with cardiac L-type VGCCs, not in a manner that conveys direct electromechanical activation, but in a way that may promote co-localization of RYR2 and the VGCC at signaling structures and potentially modulates RYR2 gating dynamics (Copello et al., 2007; Katoh et al., 2000; Mouton et al., 2001). The importance of RYR2 as a structural molecule that maintains or promotes the formation of cardiac signaling structures remains unclear. RYR2 is considered to be a protein central to cardiac function. The channel is believed to be an essential component of excitation-contraction coupling (ECC), the mechanism by which extracellular cues activated cardiomyocyte contraction (Bers, 2002). This places RYR2 at the very core of heart function. Cardiac Ca2+ signaling has also been implicated in other important processes such as setting heart rate (Monfredi et al., 2013), maintaining cardiac rhythmicity (Priori and Chen, 2011), and promoting cardiac hypertrophy and remodeling (Heineke and Molkentin, 2006). In addition, cellular Ca2+ signaling has established general roles in promoting ATP production by stimulating oxidative metabolism and in controlling cell survival and death at the level of the mitochondria (Berridge et al., 1998; Glancy and Balaban, 2012). Given RYR2’s role central role in cardiac Ca2+ signaling, it is possible that RYR2 plays an important and simultaneous role in each of these processes. As such RYR2 may have a diverse and underappreciated role in cardiac biology and disease beyond its established role in contraction signaling. The goal of this thesis is to test which specific processes are downstream of RYR2 signaling in cardiac biology.   20  1.4   RYR2 and Excitation Contraction Excitation-contraction coupling (ECC) is the mechanism by which organ level cellular signals are converted into the cellular signals which activate cardiomyocyte contraction (Bers, 2002). A heart contraction is initiated in cardiac pacemaker cells, which spontaneously depolarize, and release electrical signals that propagate rapidly throughout the heart (Monfredi et al., 2013; Rohr, 2004). Individual myocytes must respond to this electrical excitation and convert into an intracellular signal capable of activating the machinery which generates contraction force (Bers, 2002). Cardiomocytes utilize Ca2+ as the second messenger which bridges the pacemaking cue to the contractile effectors and it is thought that RYR2 plays a key role in this process. For contraction to occur, motor domains of the myosin heavy chain must be able to interact with the actin filaments and form transient cross bridges (Bers, 2002; Solaro, 2010). In the diastolic resting state, myosin is prevented from binding the actin filaments by tropomysoin, a long filamentous protein that competes for and blocks myosin binding sites on the filament (Bers, 2002; Solaro, 2010). The position of tropomyosin on actin filaments is regulated by troponin in a Ca2+ dependent manner (Bers, 2002; Davis and Tikunova, 2007; Solaro, 2010). Troponin is a complex of three proteins: troponin T, which cements troponin to tropomyosin, troponin C, which functions as a Ca2+ sensor, and troponin I, which binds actin and functions to latch the tropomysoin/troponin complex to the actin filament (Bers, 2002; Davis and Tikunova, 2007; Solaro, 2010). When Ca2+ binds troponin C, the complex undergoes an allosteric shift dislocating troponin I from actin, which causes tropomyosin to shift and uncover myosin binding sites on the actin filament (Bers, 2001; Davis and Tikunova, 2007; Solaro, 2010). Myosin can then interact with actin in a cyclical way that consumes ATP and generates force (Bers, 2002;   21 Davis and Tikunova, 2007; Solaro, 2010). This causes the thick myosin filaments to pull on the thin actin filaments, shortening sarcomeres, and causing cardiomyocyte contraction (Bers, 2002; Davis and Tikunova, 2007; Solaro, 2010). Therefore, the fundamental switch that activates heart contraction is a large (~10 µM) cytosolic Ca2+ transient which is provided by ECC and involves RYR2 (Bers, 2002; Solaro, 2010). Excitation-contraction coupling involves a variety of ion channels located in the cardomyocyte plasma membrane and SR network (Bers, 2002). To coordinate this process the heart has a specialized architecture which localizes these signaling elements to increase the efficiency of signaling (Bers and Shannon, 2013; Brette and Orchard, 2003). Cardiomyocytes are, generally speaking, cylindrical cells packed with sarcomeres that run the length of the cell (Solaro, 2010). To provide Ca2+ to core sarcomeres and to promote simultaneous signaling throughout the cell, mammalian ventricular cardiomyocytes have developed a network of invaginations known collectively as ‘t-tubules’ which have both transverse and longitudinal elements (Brette and Orchard, 2003). Wrapped around this tubule network are elements of the cardiomyocyte SR, where enlarged regions, termed the ‘junctional SR’, are enriched for SR proteins involved in ECC signaling (Bers and Shannon, 2013; Wu and Bers, 2006). At these sites, t-tubules and junctional SR form a synapse like structure called a ‘juncture’ or ‘dyad’ where plasma membrane and SR proteins involved in ECC signaling are held in close proximity to maximize their communication (Scriven et al., 2013). Specifically, this structure is important for maintaining the functional association of the cardiac L-type VGCC and RYR2: proximity is a key part of the mechanism by which relatively small Ca2+ influx via the VGCC is able to activate downstream RYR2 channels to generate the larger, global Ca2+ signals which initiate contraction (Bers, 2002; Scriven et al., 2013). This t-tubule/SR juncture is also critical for grouping other   22 proteins involved in this process including the plasma membrane NCX exchanger and SCN5 sodium channel as well as the SR SERCA2a calcium pumps which are critical for initiating and terminating ECC signals (Bers, 2002; Scriven and Moore, 2013; Scriven et al., 2013). It also bears mentioning that putative ECC signaling structures are not limited to t-tubule/SR junctions, but may also be associated with axial elements of the t-tubule network (Asghari et al., 2009) and with caveolin 3 containing regions of the plasma membrane (Schulson et al., 2011; Scriven et al., 2005). Regardless, ECC is a process highly dependent on the co-localization of signaling proteins and cellular microarchitecture.  The current model of excitation contraction is as follows (Bers, 2002). A cardiac action-potential, originating in cardiac pacemaker cells, reaches a target cardiomyocyte (Bers, 2002). This depolarizes the cardiomyocyte membrane, causing rapidly inactivating, voltage gated Na+ channels (SCN5A) located on the plasma membrane and throughout the T tubule network, to open causing a large influx of Na+ ions down its electrochemical gradient (Bers, 2002; Marban et al., 1998). Rising Na+ levels inactivates SCN5A via an inactivation gate (Marban et al., 1998). This Na+ influx further depolarizes the cell membrane causing L-type VGCC’s to open and admit a relatively small influx of Ca2+ into the cardiomyocyte cytosol at t-tubule/SR junctures (Bers, 2002; Catterall, 2011). In addition, the high Na+ levels may cause NCX channels, also localized at signaling junctures, to export Na+ and import a small amount of Ca2+ into the cell (Blaustein and Lederer, 1999; Philipson and Nicoll, 2000; Reeves and Condrescu, 2008; Scriven and Moore, 2013). The Ca2+ imported into the junctional synapse will bind to nearby RYR2 channels, causing them activate and release a much larger amount of Ca2+ into the cardiomyocyte cytosol (Bers, 2002). This serves to displace the troponin/tropomysoin complex allowing myosin to bind actin filaments and activate cellular contraction (Bers, 2002; Davis and Tikunova, 2007;   23 Solaro, 2010). RYR2 Ca2+ release will also inactivate the VGCC, which is made refractory by high cytosolic Ca2+ (via an associated calmodulin) (Peterson et al., 1999; Pitt et al., 2001; Qin et al., 1999; Zühlke et al., 1999), and cause NCX to export Ca2+ from the cell in exchange for Na+ (Bers, 2002; Blaustein and Lederer, 1999; Goldhaber and Philipson, 2012; Reeves and Condrescu, 2008). RYR2 will then inactivate and become refractory, although the exact mechanism for this remains unknown and controversial (Bers and Shannon, 2013; Guo et al., 2015; Györke and Györke, 1998; Lukyanenko et al., 1998; Sham et al., 1998; Sitsapesan and Williams, 1994). Meanwhile the overall Ca2+ signal will be terminated as the ion is cleared from the cytosol by NCX, PMCA, and SERCA2a, which will also refill the SR Ca2+ stores (Bers, 2002; Blaustein and Lederer, 1999; Brini and Carafoli, 2011; Goldhaber and Philipson, 2012; Reeves and Condrescu, 2008; Vandecaetsbeek et al., 2011). Falling Ca2+ concentration will cause the troponin/tropomysoin complex to once again block myosin binding sites on actin filaments and end contraction (Bers, 2001).  RYR2 is thought to be a critical component of excitiation-contraction coupling (Bers, 2002). Much of this is a result of decades of pharmacological studies using RYR2 inhibitors. As mentioned previously, ryanodine, an extremely high affinity and specific inhibitor of the RYR2 channel, causes a reduction in cardiac contractility and at high enough concentrations and given sufficient time (since ryanodine only binds active channels) will completely stop cardiomyocyte contraction (Jenden and Fairhurst, 1969; Meissner, 1986; Sutko et al., 1997). Similar trends are seen for other RYR2 inhibitors such as tetracaine, ruthenium red, and dantrolene (Mackrill, 2010; Thomas and Williams, 2012). RYR2 deletion is embryonic lethal at embryonic day 10, a developmental moment believed to coincide with the emergence of neonatal heart function (Takeshima et al., 1998). Lifelong Ryr2 haploinsufficiency causes mild alterations in cardiac   24 Ca2+ signaling, but does not decrease cardiac output or apparent heart contractility (Zou et al., 2011). However, one cannot rule out the possibility that this model may have substantial compensation caused by a life-long, whole body genetic manipulation (Zou et al., 2011). That said, the collective studies of RYR2 and ECC signaling provide compelling evidence for the channels involvement in this process and there is a clear consensus in the field of cardiomyocyte biology that RYR2 is critical to this process.  1.4.1   Hypothesis: RYR2 is required for cardiac function. One of the goals of this thesis is to formally test whether RYR2 is indeed critical for initiating heart contraction and cardiac function. Specifically to test the hypothesis that RYR2 is required for excitation-contraction signaling and the generation of contractile force. While there is clearly a wealth of evidence suggesting that RYR2 is important to this process, this has never been formally tested using a genetic loss of function approach in an adult organism. By using a tissue specific, inducible gene knockout approach we are able to test this hypothesis in a novel way, which could provide new insight or nuance to our current understanding of ECC signaling and heart function.  1.5   RYR2 and Heart Rate The origins of heart rate remain somewhat controversial. While there is a consensus that normal heart rate originates in specialized pacemaker cells located in the sinoatrial node of the heart, the exact proteins involved in the process and their relative contributions remain uncertain. At the time we began this project, there were essentially two camps in the field with competing, yet not   25 quite mutually exclusive theories: ‘the funny current hypothesis’ and ‘the two-clock hypothesis’ (Lakatta and DiFrancesco, 2009). What is commonly understood about heart rate is that the signal to contract normally initiates in the sinoatrial node of the heart. There, specialized cells exhibit spontaneous excitability in that their membrane potential fluctuates in cyclical, regular ways rather than resting at a stable voltage like many other excitable cell types (Monfredi et al., 2013). A key feature of this process is that during diastole, between heart contractions, pacemaker cell potential becomes highly negative and gradually depolarize by a disputed mechanism that eventually leads to activation of CaV1.2 L-type VGCCs, which rapidly fully depolarizes the cell and initiate a cardiac action-potential (DiFrancesco, 1993; Monfredi et al., 2013). The activated cardiac action potential then travels through the conductive cells of the heart and then between individual cardiomyocytes to initiate organ level contraction in a coordinated and rhythmic way (DiFrancesco, 1993; Monfredi et al., 2013; Shaw and Rudy, 1997). In this model the rate of the slow depolarization step is predicted to be the critical component of the spontaneous activity of pacemaker cells and the fundamental determinant of heart rate (DiFrancesco, 1993; Monfredi et al., 2013). It is thought that either the ‘funny current hypothesis’ or the ‘two-clock hypothesis’ explains this critical slow leak behavior (Lakatta and DiFrancesco, 2009). ‘The funny current hypothesis’ posits that heart rate is governed by the HCN family of hyperpolarization-activated, inwardly rectifying cation channels, primarily the HCN4 isoform (DiFrancesco, 2010). The HCN family of channels have several characteristics that make them attractive candidates as pacemaking channels. Several isoforms are highly expressed in SA node pacemaking cells, and they are generally enriched throughout the pacemaking and conductance systems of the heart but not seen at appreciable levels in quiescent myocytes (DiFrancesco,   26 2010; Liu et al., 2007; Shi et al., 1999). Additionally, HCN channels become maximally active at highly negative membrane potentials (-50 mV to -100 mV), such that they are able to import Na+ and K+ into cells during the electrochemical conditions of a polarized pacemaker cell, making them potentially responsible for the ‘slow phase’ of pacemaker depolarization in the SA node (Baruscotti et al., 2005; DiFrancesco, 1985). HCN channels also exhibit modification from both sympathetic and parasympathetic signals: the funny current increases in response to beta-adrenergic stimulation and is inhibited by acetylcholine in ways that are consistent with the known effects of these signals on heart rate (DiFrancesco and Tortora, 1991; DiFrancesco and Tromba, 1987; DiFrancesco, 2010). More direct evidence is that targeting deletion of cardiac HCN4 in mice results in significant bradychardia and death by atrioventricular conductance block (Baruscotti et al., 2011) and that Ivarbradine, a drug which specifically targets HCN channels, reduces heart rate (Bucchi et al., 2002; DiFrancesco and Camm, 2004). Therefore, there is compelling evidence that HCN channels generally, and HCN4 in particular, are involved in pacemaker cell biology and in setting heart rate. A number of observations have been made that run contrary to predictions made by the funny current hypothesis. For instance, while invarbardine and HCN4 deletion both decrease heart rate, neither the HCN pharmacological inhibition nor the loss of the protein abrogate the spontaneous excitability of pacemaker cells (Baruscotti et al., 2011; Bucchi et al., 2002; Denyer and Brown, 1990; DiFrancesco and Camm, 2004; Herrmann et al., 2007; Hoesl et al., 2008). Moreover, it can be argued that the death of cardiac HCN4 knockout mice is a result of dysfunction in action potential conductance rather than a consequence of reduced heart rate (Baruscotti et al., 2011). Indeed, one theory for the role of HCNs in the heart posits the channels may function as hyperpolarization brakes which prevent pacemaking and conductance cells from reaching   27 deleteriously negative membrane potentials (Herrmann et al., 2007). It has also been noted that certain vertebrate models, such as the bullfrog, display SA node dependent heart rate but have no observable ‘funny current’ in their pacemaker cells which suggests that other molecular mechanisms must exist and that HCN channels, at the very least, are not universal (Shibata and Giles, 1985). This creates the possibility that HCN channels and HCN4 may be components, albeit important ones, in a more complex pacemaking signaling system.  ‘The two-clock hypothesis’ suggests an alternate theory that pacemaking emerges from “two interlocking ionic clocks” which function together to generate the spontaneous excitability of pacemaker cells (Lakatta et al., 2010; Monfredi et al., 2013). In this theory the critical slow depolarization step requires an ensemble of ion channels, with discrete roles for proteins on both the plasma membrane and in the SR of pacemaker cells (Lakatta et al., 2010; Monfredi et al., 2013). HCN channels are still thought to contribute to the slow depolarization step, however according to this model spontaneous Ca2+ release from RYR2 channels is also a central mechanism of this process (Lakatta et al., 2010; Monfredi et al., 2013). RYR2, due to its sensitivity to luminal SR Ca2+, will experience small, local Ca2+ release events (Lakatta et al., 2010; Monfredi et al., 2013). These Ca2+ releases, which occur in very close proximity to NCX channels will cause the exchanger to export one Ca2+ ion in exchange for three Na+ ions, leading to a small net depolarization in the pacemaker cell (Lakatta et al., 2010; Monfredi et al., 2013). As the pacemaker cell depolarizes, T-type Ca2+ VGCCs, which activate at lower membrane potentials than L-type VGCCs, may also admit relatively small amounts of Ca2+ into the cell (Hagiwara et al., 1988). This will contribute to cellular depolarization and will also activate more RYR2 Ca2+ release and more NCX-dependent depolarization in a feed forward system (Lakatta et al., 2010; Monfredi et al., 2013). Recent work has additionally implicated the CaV1.3 L-type   28 VGCC, which has a lower activation voltage than CaV1.2, as having a critical role in stimulating RYR2 SR Ca2+ release during the depolarization phase (Torrente et al., 2016). Due to the combined action of HCN channels and the ongoing Ca2+ signaling being mediated primarily by RYR2 and NCX, pacemaker membrane potential will reach threshold, CaV1.2 L-type VGCCs will initiate and depolarize the cell, and a cardiac action potential will be generated (Hagiwara et al., 1988; Lakatta et al., 2010; Monfredi et al., 2013). Critically this model is also stimulated by beta-adrenergic stimulation as PKA has been shown to target RYR2, SERCA2a (via phospholamban), and L-type VGCCs to increase cardiac force and inotropy in cardiac systems (Lakatta et al., 2010; Monfredi et al., 2013; Vinogradova et al., 2006). The two-clock hypothesis then, provides a model of pacemaking with a robust and even somewhat redundant mechanism for regulating heart rate. The ‘two-clock’ hypothesis predicts that SR Ca2+ release is a critical step in pacemaking and predicts that RYR2 and NCX are essential for heart rate (Lakatta et al., 2010; Monfredi et al., 2013). Electrophysiological inhibition of NCX, by removing extracellular Na+, blocks spontaneous excitability of sinoatrial cells suggesting a non-redundant role for the protein in this process (Sanders et al., 2006). Furthermore, inducible, atrial specific NCX1 knockout in mice completely abrogates SA node pacemaking, despite normal HCN channel current and otherwise normal Ca2+ homeostasis, providing strong evidence for the ‘two-clock’ model (Groenke et al., 2013). Treatment with Ryanodine has been shown to reduce pacemaker cell slow depolarization, reduce the rate of spontaneous beating in isolated sinoatrial cells, and in many cases block the generation of cardiac action potentials outright (Ju and Allen, 1998; Li et al., 1997; Rigg et al., 2000; Satoh, 1997; Sutko and Willerson, 1980). Similarly, pharmacological inhibition of SERCA2a, which depletes the SR Ca2+ store and reduces spontaneous RYR2 release events, and   29 inhibition of NCX also cause a reduction the beating rate of sinoatrial cells in culture (Cheng et al., 2011). Stem cell derived RYR2 knockout cardiomyocytes, which model embryonic heart cells, also displayed reduced rates of spontaneous contraction which also implies that RYR2 has a role, at least in this context, in setting heart rate (Yang et al., 2002). In addition, RYR2 mutations have been found to cause catecholaminergic polymorphic ventricular tachycardia, an arrhythmic disease that can result in cardiac death (more below). While this is not necessarily evidence for RYR2 involvement in normal heart rate, it does suggest that RYR2 activity is able to activate cardiac action potentials. At the time I began this thesis, the atrial NCX study had not been published, which meant that there was a dearth of genetic, in vivo evidence for whether RYR2 and NCX were critical components of the cardiac pace-making machinery. A major prediction of the ‘two-clock’ hypothesis is that RYR2 plays a non-redundant role in setting heart rate. Another important aspect of heart rate is rhythmicity: it is critical that heart contractions occur in an orderly, coordinated manner at a rate that is consummate with the mechanical pumping action required. Arrhythmias, irregular fluctuations in the rate, can disrupt cardiopulmonary function and cause dangerous interruptions of blood flow to tissues (Goel et al., 2013). In mild cases this can result in syncopy or dizziness (Goel et al., 2013). In cases of extreme tachychardia, this can result in sudden cardiac death (Zipes and Wellens, 1998). RYR2 has been implicated as being important in cardiac rhythm through its association with the inheritable arrhythmic disease CPVT. Catecholaminergic polymorphic ventricular tachycardia (CPVT) is a rare, autosomal dominant genetic disease that causes dangerous tachycardia in response to exercise or stress (Napolitano and Priori, 2007). It effects approximately 1:10000 people worldwide and is   30 believed to account for ~15% of all unexplained cardiac deaths in young people (Napolitano and Priori, 2007). The disease is linked to an over-excitability of the heart to catecholamines, either from sympathetic enervation or from epinephrine released in the blood, which causes ventricular tachycardic arrhythmias and disrupts heart function (George, 2013; Napolitano and Priori, 2007). If this does not self-terminate, the arrhythmia can become full blown ventricular fibrillation, which in the absence of rapid medical intervention, results in sudden cardiac death (George, 2013; Napolitano and Priori, 2007).  CPVT has been mechanistically linked to RYR2 and proteins which alter the gating properties of RYR2 (Priori et al., 2001). Within RYR2, the mutations track to certain common disease ‘hot spots’ in the protein, specifically the majority of CPVT mutations are found in an n-terminal region hotspot, a central core region hotpsot, and a disease hotspot localized in the c-terminal domain (Van Petegem, 2015). The n-terminal mutations mostly correspond to an interface between RYR2 monomers in the cysotolic ‘cap’ in a region that undergoes an allosteric shift upon channel gating (Van Petegem, 2015). It has been theorized that this region may be a ‘gate’ that is involved in holding the channel closed, and that mutations in this region destabilize the closed conformation of RYR2 by disrupting interactions on this interface (Van Petegem, 2015). The exact molecular mechanism of the central domain and c-terminal domain disease hotspots remain largely unknown (Van Petegem, 2015). However, given what very recent studies have revealed about these domains and their proposed roles in Ca2+ sensing and channel gating, it is attractive to think disease mutations in these regions may alter either RYR2’s affinity for cytosolic Ca2+ or weaken this gates ability to maintain a closed conformation (Efremov et al., 2015; Yan et al., 2015; Zalk et al., 2015). CPVT mutations have also been associated with calsequestrin, triadin, and calmodulin, which are all proteins involved in modifying RYR2 gating   31 in ways that reduce open probability or increase protein refractoriness (Lahat et al., 2001; Nyegaard et al., 2012; Roux-Buisson et al., 2012). CPVT mutations have also been associated with reduced association between RYR2 and FKBP12 and FKBP12.6, which have also been shown to stabilize the closed conformation of the channel, although this remains highly controversial (Wehrens et al., 2003). Collectively though, CPVT causing mutations are considered ‘gain-of-function’, since they generally result in more permissive RYR2 activation, and therefore extra, inappropriate SR Ca2+ release (Jiang et al., 2005; 2002; Tong et al., 1997). The vast majority of functional studies have revealed that genetic knock-ins of RYR2 CPVT mutations in mice or stem cell derived cardiac cells, have born out the ‘gain-of-function’ theory since these models display increased spontaneous RYR2 opening in response to catecholaminergic stimulation (Cerrone et al., 2005; George et al., 2003; Jiang et al., 2004; 2002; 2010; Kannankeril et al., 2006; Lehnart et al., 2008; Uchinoumi et al., 2010). This has been associated with a lower activation threshold from cytosolic Ca2+, SR Ca2+, or both (George et al., 2003; Jiang et al., 2005). The current model for CPVT suggests that exercise or stress acts via beta-adrenergic stimulation and PKA to alter the collective ECC apparatus in a way that, in a system sensitized by a CPVT mutation, can cause inappropriate RYR2 Ca2+ release during diastole (Blayney and Lai, 2009; Jiang et al., 2005). It is thought that PKA activity significantly increases SR Ca2+ load by increasing Ca2+ entry into the cell via the L-type VGCC and increasing SERCA2a dependent SR Ca2+ uptake via the phospholambin regulatory protein (Blayney and Lai, 2009; Brittsan, 2000; Jiang et al., 2005). High SR Ca2+ is an activating signal for RYR2 Ca2+ release which increases RYR2’s sensitivity to cytosolic Ca2+ and can, beyond a certain threshold, directly cause RYR2 release in the absence of cytosolic Ca2+ (Blayney and Lai, 2009; Jiang et al., 2005). This is termed “store overload induced Ca2+ release” (SOICR) (Blayney   32 and Lai, 2009; Jiang et al., 2005). CPVT mutants are thought to increase RYR2 sensitivity to SOICR, by generally reducing the stability of the closed conformation of the channel, which means that RYR2 channels are more likely to open at a lower threshold of SR Ca2+ (Blayney and Lai, 2009; Jiang et al., 2005). PKA signaling, which will act to increase RYR2 open probability, further exasperates RYR2 response to SOICR in CPVT (Blayney and Lai, 2009; Jiang et al., 2005). Collectively this causes a significant number of RYR2 to release Ca2+ independently of ECC signaling, including during diastole when RYR2 are normally quiescent. Ca2+ release during diastole can cause NCX channels to export Ca2+ from the cell, which will cause a net positive charge to leak into the cell as Na+ ions are counter-exchanged (Bers et al., 2002; Blayney and Lai, 2009; Jiang et al., 2005). If this happens in sufficient quantity, it can depolarize the cell and activate voltage gated sodium channels and VGCCs initiating ECC, a cardiac action potential, and cardiac contraction inappropriately (Blayney and Lai, 2009; Jiang et al., 2005). This can then act in a feed-forward manner, where faster contractions lead to further SR Ca2+ uptake, mis-timed Ca2+ release, and erroneous contractions which results in the dangerous tachycardic incidents of CPVT (Blayney and Lai, 2009; Jiang et al., 2005). This model, the so-called ‘delayed afterdepolarization (DAD) theory’ provides a mechanism where CPVT mutations associated with reduced channel stability are able to manifest as tachycardic arrhythmias and suggests that RYR2 activity is important for cardiac rhythm (Blayney and Lai, 2009; Jiang et al., 2005).  Not every CPVT causing RYR2 mutation is associated with increased channel activity or the SOICR model of disease action. A few interesting RYR2 mutants are actually predicted to have reduced channel opening or no clear effect on single channel conductance (Jiang et al., 2007; Thomas et al., 2004; Xiao et al., 2015). One mutant is particularly interesting as it not only   33 shows slightly reduced Ca2+ release, but complete insensitivity to SOICR, meaning that it cannot cause CPVT by the DAD mechanism (Jiang et al., 2007). While the mechanism for how a reduction in RYR2 signaling can cause CPVT remains unclear, this data suggests that a reduction in RYR2 function may also be arrhythmogenic. This research also raises the possibility that appropriate, wildtype RYR2 may have a tonic role in promoting heart rhythm.  1.5.1   Hypothesis: RYR2 is required for cardiac heart rate and rhythm. One of the goals of this thesis is to formally test whether RYR2 is a key component of cardiac pacemaking. A clear prediction of the ‘two clock’ hypothesis is that RYR2 plays a critical and non-redundant role in setting heart rate in pacemaking cells. If this theory is true, deleting the Ryr2 gene in adult tissues will result in a reduction in basal heart rate. I will use my tissue specific, inducible gene knockout approach to formally test whether this prediction is true. In addition, the ‘reduced function’ CPVT mutation studies suggest RYR2 may also play a role in maintaining normal cardiac rhythm since these RYR2 mutations reveal that reducing RYR2 channel opening can cause tachycardia. Again, I will use my heart targeted, inducible gene knockout approach to study whether a reduction in RYR2 signaling will cause the emergence of cardiac arrhythmias in adult animals. This research will provide novel insight into what paces heart rate and could provide evidence that helps establish the molecular machinery responsible for cardiac pacemaking.     34 1.6   RYR2 and Cardiac Metabolism The heart is an interesting organ from a metabolic perspective. The heart, at its most basic level, is a muscular sac which pumps blood at high pressure constantly. This means the organ is constantly burning energy: generating the force of myocyte contraction and maintaining the ionic gradients needed for ECC coupling consumes a vast amount of ATP (cycling ~6kg of ATP per day in humans) (Neubauer, 2007). This means the heart must be constantly supplied with energy substrates and must maintain a very high level of ATP turnover to function. The heart is also a variably perfused organ: the force of contraction occludes the cardiac vasculature and decreases access to substrates and oxygen (Schwanke et al., 2000).  What this means is that heart metabolism must be a finely tuned balancing act that ensures energy supply is linked to demand and maintained during periods of adversely changing substrate availability. RYR2 is poised to play a critical role in this process. Given the importance of generating ATP in the heart, the organ is able to utilize a variety of energy substrates to meet its energetic demands. The heart relies on fatty acid oxidation for ~60-90% of its ATP generation, using beta-oxidation and the tricarboxylic acid (TCA) cycle to generate most of its energetic requirements (Lopaschuk et al., 2010). The remainder of cardiac ATP is generated by about equal amounts of glucose and lactate oxidation (Stanley, 2005). While glucose generates a relatively small amount of cardiac ATP, it is a significant energy substrate due to its relative oxygen efficiency (Stanley, 2005; Stanley et al., 1997). During periods of oxidative stress and hypoxia, glucose may be preferentially utilized since it requires fewer molecules of oxygen per ATP generated than fatty acid oxidation (Stanley, 2005). In addition, glycolysis generates ATP by substrate level phosphorylation, and is therefore able to generate ATP without using oxygen (Stanley, 2005). This perhaps makes glucose oxidation an   35 important fuel for periods of hypoxic stress caused by increased cardiac demand, since increased cardiac function increases the hypoxic stress on the heart. Two key takeaways from this are that TCA cycle activity is critical for effective energy production in heart and that regulation of substrate switching is important for maintaining energy production in face of changing cardiac conditions.  Mitochondrial Ca2+ is thought to promote oxidative metabolism, specifically TCA cycle activity (Glancy and Balaban, 2012). Early in vitro studies working with isolated mitochondrial enzymes revealed that Ca2+ increases the activity of a variety of TCA cycle dehydrogenases including the pyruvate dehydrogenase (PDH) complex, isocitrate dehydrogenase, and oxaloglutarate dehydrogenase (Denton et al., 1988; Denton, 2009). Other work has suggested that the electron transport chain (ETC) is also promoted by mitochondrial Ca2+ (Glancy et al., 2013). The mechanism of this remains largely unknown, but high quality data has shown that the activity of ETC complex I, complex III, and complex IV are all significantly increased by Ca2+ in isolated mitochondria (Glancy et al., 2013). The mitochondrial ATP synthase (ATPase) itself is also stimulated by mitochondrial Ca2+ (Glancy and Balaban, 2012). Again the exact mechanism of this effect is unknown, but ATPase shows a demonstrable and considerable increase in its maximum enzymatic rate (Vmax) in the presence of mitochondrial matrix Ca2+ (Glancy and Balaban, 2012). This means that mitochondrial Ca2+ entry has a role in promoting the rate at which energy substrates are converted to reducing equivalents, the efficiency of converting reducing equivalents into a proton gradient, and the speed with which this proton gradient is converted into molecules of ATP in the TCA cycle.  Pyruvate dehydrogenase complex Ca2+ sensitivity is particularly significant to cardiac metabolism. PDH is a large multi-subunit protein complex that catalyzes the conversion of   36 pyruvate to acetyl-CoA, generating NADH reducing equivalents for downstream metabolic processes (Patel et al., 2014). PDH also represents a key metabolic gatekeeper: the complex converts the final product of glycolysis into a chemical intermediary which can enter the TCA cycle for complete oxidation (Patel and Korotchkina, 2006; Randle, 1986; Sun et al., 2015). As such, PDH divides non-oxidative glucose energy metabolism from complete glucose oxidation (Patel and Korotchkina, 2006; Randle, 1986; Sun et al., 2015). Perhaps unsurprisingly, PDH activity is highly regulated as a means to control the rate of glucose oxidation in energy metabolism (Holness and Sugden, 2003; Patel and Korotchkina, 2006). PDH is regulated in part by alllostery: it is stimulated by high levels of its substrates, pyruvate, CoA, and NAD+, and inhibited by high levels of its products, Acetyl-CoA and NADH (which incidentally is a component of activating biosynthesis in the context of ‘high energy’ and the mechanism by which fatty acid oxidation interrupts glucose oxidation in the so-called ‘Randle cycle’) (Randle, 1986). PDH is also inhibited by high levels of ATP and stimulated by high ADP, meaning that the enzyme complex is also directly sensitive to energy state (Randle, 1986). In addition, PDH is regulated by an inhibitory phosphorylation event on its PDHa1 catalytic subdomain on serine 293 (Patel et al., 2014). This phosphorylation is the result of two opposing regulatory systems: the inhibitory pyruvate dehydrogenase complex kinase (PDK) family and the stimulatory pyruvate dehydrogenase complex phosphatase (PDP) family of proteins (Holness and Sugden, 2003). This regulatory phosphorylation site is also the key point of Ca2+ regulation of the PDH complex (Denton et al., 1988; Denton, 2009). PDH is therefore both a key integration site for cellular metabolic state and a major metabolic regulatory protein responsible for controlling glucose oxidation.   37 Generally, the PDKs are increased to reduce glucose oxidation in response to fatty acid availability or metabolic deficit. There are four isoforms in the pyruvate dehydrogenase kinase family of proteins (PDK1-4) (Holness and Sugden, 2003; Jeong et al., 2012; Sugden and Holness, 2006). PDK 1 and PDK 3 are transcribed by activation of hypoxia inducible factors (HIFs), which are a transcription factor system that senses both cellular O2 availability and TCA cycle state (via alpha-ketoglutarate involvement in HIF1α degradation) (Kim et al., 2006; Lu et al., 2008; Williamson et al., 1993). PDK4, the primary cardiac isoform, is generally considered a starvation and nutrition sensor since it is generated by the energy sensitive transcription factor FOXO1 (Jeong et al., 2012). PDK4 additionally is transcribed by PPARα, a master regulator of lipid metabolism, which is one of the mechanisms by which fatty acid oxidation acts to repress glucose oxidation (Huang et al., 2002; Jeong et al., 2012). The PDK family is also regulated by insulin, with PDK4, PDK1, and PDK2 being transcriptionally repressed by this hormone (Huang et al., 2002; Jeong et al., 2012). Additionally, PDK enzymatic activity is regulated by NADH, pyruvate, and ATP levels to dynamically tune their activity to cellular energy status (Pettit et al., 1975; Sugden and Holness, 2006). This provides a mechanism by which glucose availability is able to decrease repression of PDH to increase metabolic utilization of the sugar. PDKs then, integrate a wide variety of metabolic signals to provide a context sensitive break on PDH activity. PDPs, in contrast, represent a simpler regulation system. There are two isoforms in protein family, PDP1 and PDP2 (Huang et al., 1998). PDP1 has a much higher activity than PDP2, and is the dominant isoform in cardiac tissues (Huang et al., 1998). PDP1 activity is Ca2+ sensitive such that enzyme only reaches its maximal enzymatic rate in the presence of Ca2+ (Denton, 2009). The exact molecular mechanism of this effect remains controversial: there is no clear Ca2+   38 binding domain on PDP1, and it is theorized that Ca2+ may actually bind in the interface between PDP1 and the PDH complex, however the effect is well established (Denton, 2009). PDP2, conversely, does not display Ca2+ sensitivity, and is believed to provide constant low-level phosphatase activity. Since Ca2+ signaling is often involved in professional cell function, including cardiac contraction and ECC, the Ca2+ sensitivity of PDP1 represents a mechanism by which increased cellular function and energy demand can de-repress PDH and reactivate glucose oxidation. As the largest contributor to cytosolic Ca2+ in the cardiomyocyte, RYR2 is poised to interact with PDP1 and reactivate PDH function.  A major caveat of the early Ca2+ metabolic studies is that the inner mitochondrial lumen is highly impermeable to Ca2+ entry (Kohlhaas and Maack, 2013; Pozzan et al., 1994). The mitochondrial matrix, the inner-most chamber of the mitochondria and the site of the putative Ca2+ sensitive machinery important to metabolism, is primed to exclude cations which would erode the valuable charge gradient used to generate ATP (Kohlhaas and Maack, 2013). Moreover, the mitochondrial Ca2+ uniporter (MCU) the, at the time I started this thesis, theorized mitochondrial Ca2+ uptake mechanism, is quite insensitive to cytosolic Ca2+ levels, requiring a superphysiological and toxic 50 µM Ca2+ for in vitro mitochondrial Ca2+ entry (Kirichok et al., 2004). Since this level of Ca2+ does not occur in healthy functioning cells, this meant that mitochondrial Ca2+ uptake was not due to passive diffusion from the cytosolic milieu. Instead it was theorized that mitochondrial Ca2+ uptake was caused by ‘microdomain’ signaling where mitochondrial MCU are functionally coupled via high proximity to a Ca2+ release channel to drive Ca2+ uptake (Giacomello et al., 2010; Rizzuto et al., 1993). The idea of this model is that through proximity encoding, the MCU would be able to experience a very high effective Ca2+ concentration right at the site of Ca2+ release before diffusion diluted the Ca2+ signal (Giacomello   39 et al., 2010; Rizzuto et al., 1993). It is broadly analogous to how a word whispered directly into an ear is effectively louder than someone shouting in the distance. This theory has largely been born out by a body of work showing close association between mitochondria and SR/ER Ca2+ stores and release channels, the observation of discrete ‘hotspots’ of repetitive Ca2+ signalling on the inner mitochondrial membrane, and studies showing that the IP3R family of ER Ca2+ release channels stimulate mitochondrial Ca2+ uptake and oxidative metabolism in some cell types (Cárdenas et al., 2010; Dorn and Maack, 2013; Giacomello et al., 2010; Rizzuto et al., 1993). Since beginning this thesis, a renaissance of mitochondrial Ca2+ signaling has been kicked off by the discovery of the molecular identity of the MCU and some of it’s accessory proteins (MICU1, MICU2, EMRE) (Baughman et al., 2011; Csordás et al., 2013; De Stefani et al., 2011; Perocchi et al., 2010; Sancak et al., 2013). Knowledge about the molecular identity of the MCU has allowed the for direct investigation into the importance of mitochondrial uptake in mitochondrial function and confirmed that there is indeed a link between the mitochondrial uptake, the MCU, and cellular oxidative metabolism. Given the importance of oxidative metabolism and Ca2+ signalling in the heart, a variety of studies have explored the link between mitochondrial Ca2+ and energy metabolism in the heart. Work directly linking cytosolic Ca2+ signaling with PDH activation status goes back to at least 1987 when it was demonstrated that stimulated cytosolic Ca2+ increases PDH complex activation in isolated cardiomyocytes and that this effect is blocked by pharmacological inhibition of the MCU current. (Hansford, 1987). More recent work has solidified this association using modern techniques, showing that enhancing mitochondrial Ca2+ levels by inhibiting Ca2+ extrusion via the mitochondrial Na+/Ca2+ exchanger, increases the availability of NADH reducing equivalents in failing cardiomycytes (Liu and O'Rourke, 2008). At the time of starting this thesis, this was   40 largely the state of the field: cardiac mitochondrial Ca2+ had a dose dependent role on oxidative metabolism although the dynamics, molecular machinery involved, and physiological role were largely undefined. Subsequent work has implicated the importance of the cardiac SR/mitochondrial microdomain, showing that mitochondria take up Ca2+ more effectively from SR Ca2+ release than from plasma membrane NCX artificially electrophysically driven to raise cytosolic Ca2+ (Kohlhaas and Maack, 2010), and that disruption of this microdomain, through deletion of the SR/mitochondria tethering protein mitofusin 2, results in a reduction of mitochondrial Ca2+ signals and oxidative metabolism in the heart (Chen et al., 2012). Other research tracking ‘superoxide flashes’ caused as a byproduct of mitochondrial respiration, has noted that the rate of superoxide production, and therefore TCA cycle turnover, is coupled to ECC signaling on a beat-to-beat basis, showcasing a potential link between cardiac function and metabolism (Gong et al., 2014). Recent work studying MCU knockout mice have shown that lack of MCU ablates mitochondrial Ca2+ uptake in skeletal and cardiac myocytes, decreases mitochondrial oxidative metabolism, and leads to hyper-phophorylation of PDH complex in fasted mice (but not ad libidum fed mice) (Pan et al., 2013). More work, focusing on heart specific, inducible MCU gene knockout mice, revealed that mice lacking cardiac MCU showed a loss of rapid mitochondrial Ca2+ uptake and a reduced capacity to couple ATP production to cardiac function, but no change in PDH regulatory phosphorylation levels (Kwong et al., 2015). Another heart-specific MCU knockout study found that MCU knockout hearts were unable to increase energy production in response to isoproterenol stimulation of the beta-adrenergic signalling system (Luongo et al., 2015). This study also recorded increased phosphorylation of PDH in MCU knockout hearts following isoproterenol stimulation when compared to controls (Luongo et al., 2015). Deficient energy coupling to isoproterenol was also observed in a   41 dominant negative MCU study (Wu et al., 2015). Collectively, this body of work highlights the importance of cardiac mitochondrial Ca2+ uptake in oxidative metabolism and strongly implicates SR Ca2+ release in being the source of cardiac mitochondrial Ca2+. However it remains to be formally tested whether RYR2 directly plays a role in this process and which metabolic machinery is potentially downstream of RYR2 signaling.   1.6.1   Hypothesis: RYR2 is required for mitochondrial Ca2+ uptake and oxidative metabolism One of the objectives of this thesis is to test whether RYR2 is responsible for driving Ca2+ into the mitochondria and promoting energy metabolism. When I began this thesis, it was clear that mitochondrial Ca2+ homeostasis was important for ATP production in the heart, but the actual cellular machinery involved in this process was largely unknown. It is now clear that SR Ca2+ release likely drives a beat-to-beat Ca2+ transient into the mitochondria through the MCU where it generally promotes TCA cycle turnover and ATP production and may also activate the PDH complex. My work here will use my tissue specific, inducible gene knockout approach to formally test whether RYR2 is involved in Ca2+ uptake and whether RYR2 drives cardiac oxidative ATP metabolism. It will also seek to address which energy substrate utilization processes are downstream of RYR2 and whether RYR2 has a specific role in activating the PDH complex. This work will establish whether RYR2 is involved in this process and potentially solidify another molecular component of this signaling system. If RYR2 is involved in stimulating metabolic function, this work will also provide a mechanism by which cardiac function is directly coupled to energy production in the heart.    42 1.7   RYR2 and Cell Death Programmed cell death is the mechanism by which compromised cells are induced to die in a controlled manner. In multi-cellular organisms there are frequently situations where it is advantageous to kill cells that are infected with a pathogen, functionally compromised, or otherwise failing to safely remove them from the organism (Danial and Korsmeyer, 2004). There are a variety of modalities of cell death in mammalian cellular systems (Galluzzi et al., 2012). The one most commonly thought of as ‘programmed cell death’ is apoptosis, a process generally analogous to a controlled implosive demolition of a cell (Taylor et al., 2008). Key features of this process include the dissolution of the mitochondrial metabolism and proton gradient, nuclear fragmentation, and cellular blebbing (Taylor et al., 2008). Mechanistically, apoptosis involves a variety of caspase proteases, activated by a variety of cell stress sensors, which cascade to a terminal commitment step which involves ejecting mitochondrial cytochrome C, the soluble ETC electron carrier, into the cytosol, the formation of the apoptosome complex, and the activating cleavage of caspase 3, the terminal effector protease in the process (Taylor et al., 2008). Another mode of cell death is necrosis, which is generally analogous to a cellular explosion (Galluzzi et al., 2012). In many respects, necrosis is considered deleterious, an uncontrolled cellular rupture caused by a rapid, dramatic stress that kills a cell before compensatory mechanisms or apoptosis can occur (Galluzzi et al., 2012). However, in certain contexts, particularly when a cell, perhaps afflicted with a cunning virus, is able to escape apoptosis, a programmed form of necrosis (so called ‘necroptosis’) can occur (Vandenabeele et al., 2010). Key features of this process include the formation of the ‘necrosome’, a complex of RIPK1 and RIPK3 permitted by a lack of activated capsase 8, which activates ‘mixed lineage kinase domain-like protein’, MLKL, which inserts itself into lipid bilayers disrupting organelles and rupturing the plasma membrane, and   43 killing the cell (Vandenabeele et al., 2010). Another common feature of necroptosis is the formation and opening of the mitochondrial permeability transition pore (MPTP), which releases mitochondrial proteins to the cytosol and disintegrates the mitochondrial membrane potential and abolishes ATP production (Kim et al., 2003; Vandenabeele et al., 2010). Autophagy, the process by which cells digest and recycle larger structures and organelles by ‘self-eating’ is also considered a mode of cell death (Choi et al., 2013; Kroemer and Levine, 2008). Basal levels of autophagy are considered part of the normal function turnover of cells (Choi et al., 2013), however the process can be dramatically up-regulated as part of a compensatory response to cellular stress (Kroemer et al., 2010). While this is generally considered to be beneficial to cell survival, excessive and chronic autophagy can promote or even facilitate cell death (Kroemer and Levine, 2008). The exact molecular mechanism that drives autophagic cell death remain controversial, but the key features of the process are the presence and reliance of autophagic machinery to induce cell death independently of apoptosis and caspase activation (Kroemer and Levine, 2008). Cellular Ca2+ has been shown to be involved in a variety of modes of cell death, which may provide a link between RYR2 and this process in the heart. One of the main intrinsic activators of apoptotic cell death is SR/ER stress. Cells manufacture a very large amount of their total protein content within the lumen of the SR/ER, which is a constant, complex process with a certain degree of error that can result in misfolded proteins (Hetz, 2012). Accumulation of misfolded proteins in the SR/ER lumen results in activation of the ‘unfolded protein response’, a compensatory mechanism that decreases general protein synthesis, while generating alternate protein machinery intended to repair or degrade the misfolded protein and alleviate the SR/ER stress (Hetz, 2012). However, excessive or chronic SR/ER stress can initiate apoptosis (Rao et al., 2004). Two key molecular pathways associated with this are the   44 production of CHOP, a transcription factor which decreases anti-apoptotic protein Bcl-2 and increases the chances of Bax/Bak dependent cytochrome C export from the mitochondria, and the activation of Caspase 4 in an Ire1 dependent mechanism (Levine et al., 2008; Rao et al., 2004; Zinszner et al., 1998). SR/ER Ca2+ plays a critical role in SR/ER stress since the cation is an important cofactor for protein folding chaperones, such as calnexin and calreticulin, which are critical for preventing protein misfolding (Coe and Michalak, 2009; Michalak et al., 2009). Excessive SR/ER Ca2+ release or insufficient Ca2+ reuptake can ‘wash out’ the SR/ER Ca2+ store and degrade the function of folding chaperones leading to the accumulation of misfolded protein in the organelle and initiating UPR signaling (Luciani et al., 2008). RYR2 has been linked to this process in beta cells, since pharmacological inhibition of the channel (along with analogous IP3Rs) protects the cells from ER stress activated apoptosis (Luciani et al., 2008). Additionally there is evidence that SR/ER stress-induced apoptosis plays a role in cardiac disease and that beta-adrenergic inhibition, which would reduce SR Ca2+ release, has been shown to partially ameliorate this SR/ER stress induced cell death (George et al., 2011; Minamino and Kitakaze, 2010; Okada et al., 2004; Szegezdi et al., 2006).  Since RYR2 is the primary SR Ca2+ release channel in the cardiomyocyte, it is possible that the protein could be involved in SR stress dependent cell death in the heart. Ca2+ signalling has also been implicated in opening the mitochondrial permeability transition pore in both apoptosis and programmed necrosis (Tait and Green, 2010). While the molecular identity of the MPTP remains an open area of debate (Halestrap and Richardson, 2015; Morciano et al., 2015), what is accepted is that mitochondrial Ca2+ overload results in the opening of some manner of channel that spans both the inner and outer mitochondrial membranes allowing the contents of mitochondrial matrix to spill into the cytosol and completely abolish the   45 electrochemical gradient used to generate ATP (Duchen, 2000; Tait and Green, 2010). A variety of proteins have been implicated as part of the MPTP channel including the Bax/Bak complex, the voltage dependent anion carrier, cyclophilin D, and the MCU, although there is by no means conclusive evidence for the involvement of any of these components (Halestrap and Richardson, 2015; Karch et al., 2013; Morciano et al., 2015). In skeletal myocytes, cellular Ca2+ overload and excessive STIM1/Orai1 Ca2+ uptake is associated with cellular death in a MPTP dependent manner while SERCA1 overexpression has been shown to ameliorate cellular Ca2+ levels and reduce MPTP opening (Goonasekera et al., 2014; 2011; Mázala et al., 2015). Moreover, the very recent work done on both the whole body and heart specific MCU knockout mice reveal that MPTP opening is abrogated when rapid mitochondrial Ca2+ uptake is blocked by deletion of MCU (Kwong et al., 2015; Luongo et al., 2015; Pan et al., 2013). Given RYR2s potential role as a driver of MCU Ca2+ uptake (discussed above), it is feasible that the protein may have a critical role in mitochondrial Ca2+ overload and MPTP opening in cardiomyocytes. Indeed, work in embryonic cardiomyocyte models has demonstrated that splice variants of RYR2 are protected from caffeine induced apoptosis via BCL-2/BAX/BAK mitochondrial mechanism (George et al., 2007b), which is implicated in apoptosis (Reed, 1994) and MPTP opening (Karch et al., 2013). However, a role for RYR2 in this process in adult tissues remains to be elucidated. While excessive Ca2+ entry into the mitochondria has been implicated in cell death, the converse, too little mitochondrial Ca2+ entry, has also been associated with programmed cell death. Specifically, research conducted on pancreatic beta cells revealed that pharmacological blockage of RYR2 leads to auto-activation of calpain 10, an atypical Ca2+ sensitive protease, which leads to cell death (Johnson, 2004). Calpain 10, a protein originally discovered in a human population with a very high incidence of type 2 diabetes (Horikawa et al., 2000), is a   46 mitochondrial resident protease that undergoes self-cleavage and activation due to changes in Ca2+ levels via a poorly understood mechanism (Arrington et al., 2006; Ma et al., 2001). When mitochondrial Ca2+ falls below a certain threshold, functional data suggests this protein self cleaves, releasing an active protease that targets various mitochondrial proteins and initiates cell death (Johnson, 2004). Some interesting features of this study are that the RYR2 inhibition cell death was not associated with apoptosis, was abrogated on a calpain 10 knockout background, and was only observed when beta cells were cultured in low glucose media (presumably because this also reduced IP3R signaling, which likely plays a degenerate role in mitochondrial Ca2+ homeostasis in beta cells) (Johnson, 2004). What this study overall implies is that a tonic level of Ca2+ entry into the mitochondrial matrix is a pro-survival signal and that RYR2 can be one of the proteins involved in this process (Johnson, 2004). Given that RYR2 is the dominant SR/ER Ca2+ release channel and that it has a putative role as an MCU signaling partner, it is possible that the channel plays a role in inhibiting calpain 10 activation in the heart as well.  1.7.1   Hypothesis: RYR2 is required for cell survival. One of the objectives of this thesis is to test whether RYR2 is involved in cardiomyocyte cell death and to determine which cell death modality, if any, the protein is involved with. I will therefore use my inducible, cardiac specific knockout model to test whether RYR2 knockout hearts display an altered level of cell death. I will also probe for key effectors involved in Ca2+ dependent cell death including active caspase 3, the induction of CHOP, and activated calpain 10. This work will provide evidence potentially linking RYR2 to cardiac cell death and may provide evidence for a to date unappreciated cell death mechanism in the heart    47 1.8   RYR2 and Cardiac Hypertrophy The heart is a dynamic organ that must constantly adapt to changing respiratory demand and hemodynamic conditions to ensure survival. Short-term changes in respiratory demand are accounted for by changes in heart rate and by the inherent capacity of sarcomeres to vary their contractile force in response to cardiac stretch (the so-called ‘starling force’ effect (Konhilas et al., 2002)). Longer term increases in cardiac demand (or alternately deficits in the heart’s capacity to satisfy demand), often through athletic training or pathophysiology, cause the heart and cardiomyocytes to increase in size and hypertrophy to increase the capacity and force generating potential of the organ. There are a variety of transcriptional mechanisms implicated in physiological and pathological hypertrophy, but many involve Ca2+ signals as key effectors of the process. Generally speaking there are three main Ca2+ sensitive hypertrophic signalling cascades considered significant in the heart (Heineke and Molkentin, 2006). The first is the calcinuerin/NFAT pathway (Molkentin, 2004; Wilkins et al., 2004). Prolonged increases in cytosolic Ca2+ activate the Ca2+/calmodulin dependent phosphatase, which will dephosphorylate cytosolic NFAT (Molkentin, 2004; Wilkins et al., 2004). Dephosphorylated NFAT can then migrate into the nucleus, bind to its target DNA, and promote the transcription of hypertrophic effector genes including transcription factors GATA4 and GATA6 (Molkentin, 2004; Wilkins et al., 2004). Ca2+ signals may also activate a CaMK dependent mechanism where increases in nuclear Ca2+ may activate a pool of CaMKII or CaMKIV which may then phosphorylate HDAC4, causing the histone condensing protein to be exported from the nucleus (Passier et al., 2000; Zhang et al., 2002). This can de-repress the transcription of a number of genes, including the transcription factor MEF2, which can then promote a hypertrophic gene program (Passier et   48 al., 2000; Zhang et al., 2002). Other research has implicated an ERK/MAPK dependent pathway, where increased cytosolic Ca2+ leads to the phosphorylation of ERK via a Ras/Raf dependent mechanism (Molkentin, 2004; Yamaguchi et al., 2011). pERK can activate the p90RSK kinase, which can then act to repress GSK-3β (Molkentin, 2004; Yamaguchi et al., 2011). GSK-3β normally acts to repress the transcription of NFAT and GATA4, and repression of this molecule allows these transcription factors to activate a hypertrophic response (Molkentin, 2004; Yamaguchi et al., 2011). While all three of these responses have been implicated in cardiac hypertrophy, the significance, relative contribution, context of activation, and upstream Ca2+ sources remain an area of ongoing research. A variety of Ca2+ channels have been implicated in responding to extracellular hypertrophic signaling cues. Angiotensin II, a pro-hypertrophic hormone activated in response to hypertension as part of the renin-angiotensin system, has been shown to act via a G-protein coupled receptor to activate phospholipase C, which in turn, generates DAG and IP3 (Wu et al., 2010). DAG can activate the TRPC family of plasma membrane Ca2+ channels (specifically TRPC3/6/7) to increase cytosolic Ca2+ enough to activate hypertrophic signaling via the calcineurin/NFAT pathway (Eder and Molkentin, 2011; Wu et al., 2010). There is also evidence that TRPC1/4/5 may be involved, although its unclear if this is because they form heterotrimers with DAG sensitive TRPC3/6/7 or because they have also been activated due to increased cardiac stretch (Eder and Molkentin, 2011; Wu et al., 2010). The IP3 generated from this process can also activate a pool of perinuclear IP3R2, which may act to elevate nuclear Ca2+ independently of cytosolic Ca2+ to promote hypertrophy via the calcineurin/NFAT pathway, CaMK/HDAC/MEF2 pathway, or both (Arantes et al., 2012; Nakayama et al., 2010). Additionally, it has been shown that L-type Ca2+ signaling is a requirement in some forms of cardiac hypertrophy, including   49 TRPC-dependent activation, implying that normal ECC Ca2+ homeostasis may be involved in the process as well (Gao et al., 2012). Yet other research has shown that increasing store-operated Ca2+ entry via STIM1/Orai1 Ca2+ uptake can also cause cardiac hypertrophy (Voelkers et al., 2010). Collectively this research cements the importance of Ca2+ channels in the hypertrophic process, but also raises the question of whether hypertrophic gene programs activate in response to discrete signaling cascades or in response to a general increases in the collective cellular Ca2+ level.  If hypertrophy is dependent on the overall Ca2+ temperature of a cell, than RYR2, the largest contributor to cytosolic Ca2+ transients in cardiomyocytes, may be involved in hypertrophic signalling. A number of studies have attempted to address this question. One study, utilizing whole body, constitutive RYR2 haploinsufficiency demonstrated that RYR2-heterozygous hearts show less calcineurin/NFAT dependent hypertrophy in response to aortic banding induced pressure overload, implying that RYR2 normally contributes to this pathway (Zou et al., 2011). Another study modeled increased RYR2 leak by mutating the calmodulin binding site on RYR2, thus removing an inhibitory signal and deleteriously increasing RYR2 Ca2+ release (Yamaguchi et al., 2011). This study found this pathophysiological increase in RYR2 signaling lead to cardiac hypertrophy that was not dependent on calcineurin/NFAT or CaMK/HDAC/MEF2 signaling, but was instead caused by the ERK dependent activation of NFAT and GATA4 (Yamaguchi et al., 2011). Both of these studies reveal that RYR2 plays an important role in cardiac hypertrophy but leave it an open question which hypertrophic pathways are downstream of RYR2 signaling.    50 1.8.1   Hypothesis: RYR2 is required for cardiac hypertrophy One of the minor objectives of this thesis is to observe if RYR2 is required for cardiac hypertrophy. If, as predicted, RYR2 is a major contributor to cardiac hypertrophy or if an increase in overall Ca2+ levels is necessary for the induction of hypertrophic gene programming, then deletion of Ryr2 in vivo should demonstrate reduced hypertrophy. In this case I will simply observe whether or not hypertrophy is seen in Ryr2 KO hearts.  1.9   RYR2 and Heart Disease Heart disease and failure remain one of the leading causes of death in the developed world. While modern advances in surgical intervention and drug treatment regimes have significantly improved the prognosis of people afflicted with heart disease, heart failure still affects approximately 500000 Canadians and millions of people worldwide (Blair et al., 2013; McMurray and Pfeffer, 2005). Moreover, since heart disease and failure are common complications of type II diabetes, the incidence of heart failure may increase as the incidence of obesity and metabolic disease worldwide continues to climb (Blair et al., 2013; McMurray and Pfeffer, 2005). It therefore remains critical to understand the molecular mechanisms contributing to cardiac pathology and to differentiate different modalities of heart disease. RYR2 dysfunction has been implicated in a variety of models of heart disease. Reduced RYR2 expression, density, or function has been observed in a variety of animal models of hypertensive heart disease caused by aortic constriction (Brillantes et al., 1992; Matsui et al., 1995; Milnes, 2001; Naudin et al., 1991). In addition, reductions in channel expression and function, even up to a 50% decrease, has been seen in animal models of diabetic cardiomyopathy  (Bidasee et al., 2001; Dincer et al., 2006; Shao et al., 2007; 2012; Yu et al., 1994).  Other studies,   51 using an aortic constriction model or clinical sections, showed that RYR2 localization was greatly altered in heart failure, enriching along the longitudinal elements of the t-tubule network (Crossman et al., 2011; Wu et al., 2012). Yet other studies have also observed that RYR2 expression is decreased as a normal consequence of aging, which may provide a mechanistic link between RYR2 and the progressive decrease in heart function in the elderly 2 (Assayag et al., 1998; Kandilci et al., 2011; Tellez et al., 2011). Ryr2 genetic polymorphisms have also been causally linked with hypertension in an unbiased GWAS study, although whether changes in cardiac RYR2 play a causal role remains undetermined (Wellcome Trust Case Control Consortium, 2007). While there is no clear consensus that RYR2 dysfunction does drive cardiac pathology in humans, these studies at least suggest that the idea is worthy of consideration. What is more commonly accepted though, is that SR Ca2+ release is reduced as a consequence of heart failure (Balke and Shorofsky, 1998; Bers, 2006; Gomez, 1997).  It is a known feature of late stage heart failure that SR Ca2+ stores become gradually depleted and that this results in less SR Ca2+ release during ECC and an increased reliance on plasma membrane Ca2+ dynamics to support cardiac contraction (Balke and Shorofsky, 1998; Bers, 2006; Gomez, 1997). It is generally believed that this is a result of increased Ca2+ leak from the SR and reduced Ca2+ reuptake as a result of chronic beta adrenergic signaling during pathology and increased expression of NCX exchangers during heart failure (Balke and Shorofsky, 1998; Bers, 2006; Gomez, 1997). A consequence of this, though, is that RYR2 Ca2+ release is substantially decreased in late stage heart failure regardless of changes in channel function or gating.  Dilated Cardiomyopathy, which is caused by a variety of insults including chronic hypertension, diabetes, and mutations in contractile machinery and Ca2+ handling proteins, is a multi-faceted disease with many coincident changes in cardiac function (Jefferies and Towbin,   52 2010). As the name suggests DCM is associated with cardiac hypertrophy and fibrosis which results in enlarged heart chambers, but without significantly enlarged chamber walls, as well as a reduction in the contractility and force generating potential of the organ (Jefferies and Towbin, 2010). DCM is also associated with alterations in heart rate including a general reduction in basal heart rate and the generation of atrial fibrillations and arrhythmias. Heart failure generally is also associated with metabolic abnormalities including a general reduction in oxidative energy metabolism and an increased reliance in glycolysis for ATP generation (Jefferies and Towbin, 2010). Diabetic cardiomyopathy is associated with a sharp decrease in the hearts capacity to utilize glucose as an energy substrate, and an increased reliance on FA oxidation for energy (Lopaschuk, 2002; Schilling and Mann, 2012). Given that RYR2 (as discussed above) is potentially involved in hypertrophic signaling, cardiac contraction, heart rate and rhythmicity, as well as regulating oxidative metabolism, it is possible that dysfunction or reduced signaling from this Ca2+ channel may contribute or cause some aspects of cardiac pathophysiology.  1.9.1   Hypothesis: Decreases in Ryr2 expression cause dilated cardiomyopathy  Another minor objective of this thesis is to test whether decreasing RYR2 expression and therefore function is able to cause heart failure and hypertrophy. I will use my inducible tissue specific Ryr2 knockout model to test whether an uncontrolled decrease in RYR2 levels and function can cause heart failure and observe which facets of DCM are recapitulated. I will also utilize an inducible, tissue specific Ryr2 haploinsufficiency model where I knock out one allele of the protein to test whether a stable decrease in RYR2 levels, similar to what is seen in some models of heart disease, is also able to cause heart failure. This work will formally test whether   53 RYR2 dysfunction is an upstream driver of heart failure and may elucidate the channel’s validity as a potential therapeutic target.      54 Chapter 2:  Materials and Methods  2.1   Experimental Animals Mice bearing the Ryr2 floxed allele were generated using R1 embryonic stem cells (129SvJ x 129Sv) containing a Ryr2 floxed allele (Yang et al., 2002), injected into C57BL/6 blastocysts and transferred to pseudopregnant females at the NIA transgenic facility. Agouti mice were identified and genotyping confirmed transmission of the floxed allele to offspring, which were backcrossed over 10 generations to the C57BL/6 background. Tamoxifen-inducible, cardiomycyte-specific Ryr2 knockout mice were generated by crossing the mice harboring conditional Ryr2 alleles (C57Bl6 Ryr2flox/flox mice) with mice expressing inducible Cre-recombinase under the control of the α-MHC promoter (C57Bl6-mhy6-mer-Cre-mer mice) (Sohal et al., 2001). Tamoxifen was injected into the intraperitoneal cavity of 8-20 week-old Ryr2flox/flox:mhy6-mer-Cre-mer mice (referred to as cRyr2KO mice) or littermate controls for three consecutive days at 3 mg per 40 g body weight. It is important to note that since both groups received tamoxifen, none of the effects reported herein can be attributed to this drug.  Tamoxifen-inducible, cardiomyocyte-specific Ryr2 haploinsufficiency mice (cRyr2Δ50 mice) were generated by crossing mice harboring Ryr2 alleles containing flanking loxP sites (C57Bl6 Ryr2flox/wildtype mice) with mice expressing inducible Cre-recombinase under the control of the α-myosin heavy chain promoter (C57Bl6 mer-Cre-mer mice) (Strain: 005657, The Jackson Laboratory, Bar Harbor, ME, USA). Tamoxifen was injected into the intraperitoneal cavity of 8-16 week-old Ryr2flox/wildtype:mer-Cre-mer mice and littermate “control” mice (Ryr2flox/wildtype mice injected with tamoxifen) for three consecutive days at 3 mg per 40 g of body   55 weight. All mice were given at least 3 weeks recovery time after tamoxifen treatment to reach a stable level of RYR2 ablation and to circumvent any potential effects of tamoxifen which is cleared from mice within 21 days (Koitabashi et al., 2009).  C57BI6 SILAM mice (Zanivan et al., 2012) were generated by feeding mice for two generations the MouseExpress L-LYSINE (13C6, 99%) 8g/kg diet (Cambridge Isotope Laboratories, Andover, MA, USA). Amino acid incorporation within long-lived neuronal tissue was checked and confirmed to be >97%.   All animal protocols were approved by the University of British Columbia Animal Care Committee in accordance with international guidelines.  2.2   In Vivo Analysis of Cardiac Function and Heart Rate Cardiac function was examined by echocardiography utilizing a Vevo 770 high-resolution image system (Viewsonics, Brea, CA, USA) (Gomez and Richard, 2004). Images were obtained through the anterior and posterior left ventricular walls at the papillary muscle level. Left ventricular mass and systolic function, left ventricular end-systolic and end-diastolic dimensions, interventricular septum thickness and posterior wall were measured from M-mode traces. Shortening and ejection fraction were calculated as described (Gomez and Richard, 2004). We assessed heart rate before and after tamoxifen injection. Mice assessed by echocardiography were anaesthetized with isoflurane at the minimum gas concentration necessary to maintain light anaesthesia.  In vivo heart rate and ECG was assessed utilizing ETA-F10 implantable telemetry devices according to the manufacturer’s instructions (Data Sciences International, St, Paul, MN, USA).   56 Inhaled isoflurane was used as an anesthetic during implantation surgery and injected analgesic, Buprenorphine (0.1 mg/kg), was administered daily as necessary. Heart rate and rhythmicity were analyzed following recovery from surgery and after the discontinuation of all analgesics.   2.3   Ex Vivo Analysis of Cardiac Function and Metabolism Heart function was further measured using the working heart model as described (Wambolt et al., 2006). Peak systolic pressure, aortic flow, cardiac output, and work were analyzed as previously described (Allard et al., 2007). These mice were euthanized by having their hearts excised under a dose of inhaled isoflurane sufficient to maintain deep anaesthesia. Myocardial substrate utilization was measured in working hearts, as detailed elsewhere (Allard et al., 1994; Belke et al., 1999; Khairallah et al., 2004). Briefly, working hearts were perfused with Krebs-Henseleit solution containing 11 mM glucose, 1.5 mM lactate, 0.15 mM pyruvate, 0.6 mM palmitate and 20 mU/ml insulin in a recirculating mode. Glycolysis, as well as myocardial rates of oxidation of palmitate, glucose, and lactate were determined by the quantitative collection of 3H2O or 14CO2 produced by hearts perfused with either KH solution containing [9,10-3H]-palmitate and [U-14C]-lactate or [U-14C]-glucose and 5-[3H]glucose.  2.4   Light and Electron Microscopy in the cRyr2KO Model Paraformaldehyde-fixed ventricular tissue sections were stained with H&E and Masson’s trichrome to assess fibrosis. Apoptosis and cell death of adult ventricular myocytes was assessed in heart tissue sections using the terminal deoxynucleotidyl transferase biotin-dUTP nick end-labeling (TUNEL) assay from Roche (Mississauga, Canada) according to the manufacturer’s   57 instructions. Images were taken with a Retiga-2000R camera (Q-Imaging). TUNEL-positive cells were normalized to tissue area. Live-cell imaging was conducted on cardiomyocytes isolated from cRyr2KO and control mice using Langendorff reverse perfusion to introduce collagenase to the cardiac vasculature as detailed elsewhere (Fernando, 2006). To assess cytosolic Ca2+ levels, isolated cardiomyocytes were loaded with 5 µM Fura-2-AM (Invitrogen) for 30 minutes and washed twice for 15 minutes before imaging. Fura-2-AM loaded cells were excited at 340 nm and 380 nm and measured for emission at 510 nm using a imaging system equipped with a 10x air objective (0.3 NA) on a Zeiss Axiovert-200M microscope with a CoolSnapHQ2 CCD Camera (Intelligent Imaging Innovations, Denver, CO) at 10x magnification. Ratiometric images were quantified using Slidebook software (Intelligent Imaging Innovations) to measure relative cytosolic Ca2+ (Luciani et al., 2008). To assess mitochondrial Ca2+ levels isolated cardiomyocytes were loaded with 5 µM Rhod-2-AM (Invitrogen) for 30 minutes and washed for 15 minutes before imaging. In some studies, 5 µM Mitotracker Deep Red (FM) (Invitrogen) was co-loaded. Loaded cells were imaged at the recommended wavelengths using the imaging Axiovert-200M described above and 40x oil objective (1.3 NA). Rhod-2 fluorescence was used to measure relative mitochondrial Ca2+ (Trollinger et al., 1997) and Mitotracker was used to assess relative mitochondrial content for normalization. The co-loading of these dyes did not alter Ca2+ or cell viability (not shown). All live-cell microscopy occurred within 8 hours of isolation and only rod shaped cardiomyocytes with typical cell morphology were considered in analysis.    58 2.5   Cardiomyocyte Ca2+ and Functional Measurements in the cRyr2Δ50 Model Live cell analysis was conducted on cardiomyocytes isolated from cRyr2Δ50 and control mice using Langendorff reverse perfusion to introduce collagenase in to the cardiac vasculature as detailed elsewhere (Fernando, 2006). Cardiomyocyte function and contractility were assessed as described elsewhere (Williams et al., 2014). Briefly, a suspension of the isolated ventricular cardiomyocytes was transferred to a chamber with stimulating electrodes (Cell MicroControls, Norfolk, VA, USA) that was precoated each day with laminin (1 mg/ml) to help immobilize the cells, and was fixed to the heated stage of an Olympus IX70 inverted microscope with 400X quartz optics. The cells were continuously superfused with Tyrode solution consisting of 1.5 mmol/l CaCl2 in 95% O2-5% CO2 at 0.5–1 ml/min at 34-36 °C. An IonOptix (Milton, MA, USA) video system imaged the cells at 240 hz allowing measurement of the degree and rate of myocyte shortening during field stimulation at various frequencies. To measure Ca2+ signals, we employed a ratiometric Ca2+-sensitive fluorescent dye, Fura-2-acetoxymethyester (AM) (Invitrogen, Carlsbad, CA, USA) which will localize in the cytosol, and a Ca2+-sensitive fluorescent dye that localizes in the mitochondria, Rhod-2-AM (Invitrogen, Carlsbad, CA, USA). Mitochondrial Rhod-2-AM localization was confirmed by co-loading cardiomyocytes with 5 µM Mitotracker Deep Red (644/655 nm) (Invitrogen, Calsbad, CA, USA) and imaging at high resolution using a 100x 1.45 NA objective on a Zeiss Axiovert-200 M microscope with a CoolSnapHQ2 CCD Camera  (Intelligent Imaging Innovations, Denver, CO, USA). These image stacks were deconvolved using Slidebook software (Intelligent Imaging Innovations, Denver, CO, USA). Simultaneous cytosolic Ca2+ and mitochondrial Ca2+ measurements were performed on isolated cardiomyocytes which were incubated with 5 µM Fura-2-AM (340 nm and 380 nm excitation; 510 nm emission) and 5 µM Rhod-2-AM (550 nm   59 excitation /580 nm emission) for 15 minutes and washed for 5 min before imaging using a 10x 0.5 NA objective Zeiss Axiovert-200 M microscope with a CoolSnapHQ2 CCD Camera (Intelligent Imaging Innovations, Denver, CO, USA). Additionally, imaged cells were provided field stimulation of 80 mA with a hybrid system using a stimulus isolator (World Precision Instruments, Sarasota, CA, USA) and a custom-made programmable stimulator (McAfee Scientific, Port Hardy, WA, USA) with periods of: individual analog pulses, 2 Hz continuous stimulation, or 6 Hz continuous stimulation. Fura-2 ratios to measure relative cytosolic Ca2+ and Rhod-2 intensities to measure mitochondrial Ca2+ were quantified using Slidebook software (Intelligent Imaging Innovations, Denver, CO, USA). Live-cell imaging occurred within 8 hours of cell isolation, and only rod-shaped cardiomyocytes with typical cell morphology were considered in analysis. Additionally, only cardiomycytes that displayed stable un-stimulated Fura-2 and Rhod-2 baselines as well as excitability to initial 5 field stimulation pulses were considered in the analysis. Cardiomyocytes from at least 3 independent isolations per treatment group were studied.  2.6   Gene and Protein Expression Analysis  RNA was isolated from heart tissue using Trizol, followed by cleanup using the RNeasy kit (Qiagen). After reverse transcription (SuperScript III; Invitrogen), TaqMan quantitative RT-PCR (qPCR) was conducted using probes from Applied Biosystems and PerfeCTa qPCR SuperMix (Quanta) on a StepOnePlus thermocycler (Applied Biosystems). SYBR Green quantitative RT-PCR was conducted using PerfeCTa SYBR Green qPCR SuperMix (Quanta). Relative gene expression changes were analyzed by the 2-ΔCt method and plotted in normalized relative units (RU). HPRT and cyclophilin were used as internal controls, after ensuring that they were not   60 altered in cRyr2KO cardiomyocytes. Primer details are available in Tables 2.1-2.3. Immunoblots were performed on lysates from mechanically disrupted heart, homogenized and sonicated in ice-cold lysis buffer. Samples were quantified, boiled with loading dye, and 15-100 µg of protein were used in SDS-PAGE electrophoresis. Proteins were then transferred to PVDF membranes using standard semi-dry transfer (for protein smaller than 120 kDa) or wet transfer (for proteins larger than 120 kDa), and subsequently treated with targeted primary and horseradish peroxidase-conjugated secondary antibodies. Bands were visualized using an enhanced chemiluminescence detection kit, and quantified by densitometry. Anti-HSP90 (Cell Signalling) and anti-EEA1 (Abcam) was used as loading controls for high kDa immunoblots. Rabbit polyclonal anti-RYR2 antibodies were provided by Dr. Anthony Lai (Priori and Chen, 2011). Commercially available antibody details can be found in Table S4. 	  	  	  	  	  	  	  	  	  	  	  	  Table 2.1. Sybr Green RT-qPCR probes  Gene	   Forward	  Primer	   Reverse	  Primer	  Anp	   5’-­‐TACAGTGCGGTGTCCAACACAG-­‐3’	   5’-­‐TGCTTCCTCAGTCTGCTCACTC-­‐3’	  Bnp	   5’-­‐TCCTAGCCAGTCTCCAGAGCAA-­‐3’	   5’-­‐	  GGTCCTTCAAGAGCTGTCTCTG-­‐3’	  Mhy6a	   5’-­‐GCTGGAAGATGAGTGCTCAGAG-­‐3’	   5’-­‐	  CCAGCCATCTCCTCTGTTAGGT-­‐3’	  Mhy7a	   5’-­‐GCTGGAAGATGAGTGCTCAGAG-­‐3’	   5’	  -­‐TCCAAACCAGCCATCTCCTCTG-­‐3’	  Atf4	   5’-­‐ATGGCCGGCTATGGATGAT-­‐3’	   5’-­‐CGAAGTCAAACTCTTTCAGATCCATT-­‐3’	  Bip	   5’-­‐GGTGCAGCAGGACATCAAGTT-­‐3’	   5’-­‐CCCACCTCCAATATCAACTTGA-­‐3’	  Chop	   5’-­‐CTGCCTTTCACCTTGGAGAC-­‐3’	   5’-­‐CGTTTCCTGGGGATGAGATA-­‐3’	  Gadd34	   5’-­‐CCCGAGATTCCTCTAAAAGC-­‐3’	   5’-­‐CCAGACAGCAAGGAAATGG-­‐3’	  sXbp1	   5’-­‐GAGTCCGCAGCAGGTG-­‐3’	   5’-­‐GTGTCAGAGTCCATGGGA-­‐3’	  Bax	   5’-­‐GGAGCAGCTTGGGAGCG-­‐3’	   5’-­‐AAAAGGCCCCTGTCTTCATGA-­‐3’	  PDK1	   5’-­‐	  TGGACTTCGGGTCAGTGAAT-­‐3’	   5’-­‐	  GGCTTTGGATATACCAACTTTGCA-­‐3’	  PDK2	   5’-­‐CAACACACCCTCATCTTTGATGG-­‐3’	   5’-­‐	  ATGTCATAGGCGTCTTTCACCA-­‐3’	  PDK3	   5’-­‐	  GACTTCGGAAGGGATAATGCATG-­‐3’	   5’-­‐	  TCTGCATGTACCAGCTCTGAA-­‐3’	  PDK4	   5’-­‐	  CCTGTCAGAGTTTGTAGACACG-­‐3’	   5’-­‐	  CACTGAATATGAGGATGTGCTGATTC-­‐3’	  Cyc	   5’-­‐GTCTGCAAACAGCTCGAA-­‐3’	   5’-­‐ACGCCACTGTCGCTTT-­‐3’	  Hprt1	   5’-­‐TCCTCCTCAGACCGCTTTT-­‐3’	   5’-­‐CCTGGTTCATCGCTAATC-­‐3’	  Primers	  produced	  by	  Integrated	  DNA	  Technologies	    61    	  	  Gene	   Catalogue	  No.	   Vendor	   Gene	   Catalogue	  No.	   Vendor	  Ryr2	   Mm00465914_m1	   ABI	   Fabp4	   Mm01295675_g1	   ABI	  Ryr1	   Mm01175211_m1	   ABI	   Glut4	   Mm01245502_m1	   ABI	  Ryr3	   Mm01328416_m1	   ABI	   Cd36	   Mm01135198_m1	   ABI	  Ip3r1	   Mm01183029_m1	   ABI	   Pck1	   Mm01247058_m1	   ABI	  Ip3r2	   Mm01255198_m1	   ABI	   Ppara	   Mm00440939_m1	   ABI	  Ip3r3	   Mm01306099_m1	   ABI	   Pparag	   Mm00440945_m1	   ABI	  Insr	   Mm01211875_m1	   ABI	   Ppargc1a	   Mm00447181_m1	   ABI	  Ptpn1	   Mm00448427_m1	   ABI	   Srebp1c	   Mm01138344_m1	   ABI	  Bcl2	   Mm00477631_m1	   ABI	   Chrebp	   Mm00498811_m1	   ABI	  BclX	   Mm00437783_m1	   ABI	   Foxa2	   Mm00839704_mH	   ABI	  Ucp1	   Mm01244861_m1	   ABI	   Foxc2	   Mm00546194_s1	   ABI	  Ucp2	   Mm00627599_m1	   ABI	   Foxo1	   Mm00490672_m1	   ABI	  Ucp3	   Mm00494077_m1	   ABI	   Lxr	   Mm00443454_m1	   ABI	  Acly	   Mm00652520_m1	   ABI	   Egr1	   Mm00656724_m1	   ABI	  Hsl	   Mm00495359_m1	   ABI	   Egr2	   Mm00456650_m1	   ABI	  Fasn	   Mm00662319_m1	   ABI	   Atf3	   Mm00476032_m1	   ABI	  Adpn	   Mm00504420_m1	   ABI	   Acacb	   Mm01204683_m1	   ABI	  Atgl	   Mm00503040_m1	   ABI	   Scd1	   Mm01197142_m1	   ABI	  IL6	   Mm99999064_m1	   ABI	   Adipor1	   Mm01291331_m1	   ABI	  Tnf	   Mm00443258_m1	   ABI	   Adipor2	   Mm01184030_m1	   ABI	  Ikbkb	   Mm01222244_m1	   ABI	   Adipoq	   Mm00456425_m1	   ABI	  Cebpa	   Mm01265914_s1	   ABI	   Emr1	   Mm00802530_m1	   ABI	  Hprt1	   Mm00446968_m1	   ABI	   Klf15	   Mm00517792_m1	   ABI	  ABI=Applied	  Biosystems	  Inc.	  	  Table 2.2. Commercially available Taqman RT-qPCR assays. 	  	  	  	  Gene	   Forward	  Primer	   Reverse	  Primer	   Probe	  Cyc	   5’-­‐GTCTGCAAACAGCTCGAA-­‐3’	   5’-­‐ACGCCACTGTCGCTTT-­‐3’	   5’-­‐/56-­‐FAM/TGCAGCCATGGTCAAC-­‐3’	  Produced	  by	  Integrated	  DNA	  technologies	  	  Table 2.3. Custom Taqman RT-qPCR assays.   62 	  	  	  	  	  	  	  Gene	   Catalogue	  Number	   Vendor	   Antibody	  Conce.	   Protein	  Load	  Pres1	   #3622	   Cell	  Signaling	   1/500	   75µg	  Pres2	   #2192	   Cell	  Signaling	   1/1000	   50µg	  Cl.	  Casp3	   #9664	   Cell	  Signaling	   1/1000	   15µg	  CHOP	   sc-­‐575	   Santa	  Cruz	   1/1000	   15µg	  Calpain10	   ab10820	   Abcam	   1/1000	   15µg	  Calpain1	   ab16680	   Abcam	   1/1000	   15µg	  Calpain2	   ab16261	   Abcam	   1/1000	   15µg	  INSR	   #3020	   Cell	  Signaling	   1/1000	   50µg	  pAktT308	   #4056	   Cell	  Signaling	   1/1000	   15µg	  pAktS473	   #9271	   Cell	  Signaling	   1/1000	   15µg	  Akt	   #9272	   Cell	  Signaling	   1/1000	   15µg	  pERK	   #9106	   Cell	  Signaling	   1/1000	   15µg	  ERK	   #9102	   Cell	  Signaling	   1/1000	   15µg	  HIF1α	   NB	  100-­‐123	   Novus	  Biologicals	   1/1000	   50µg	  HIF1β	   611078	   BD	  Biosciences	   1/1000	   50µg	  LC3	   0260-­‐100/LC3-­‐2G6	   Nanotools	   1/1000	   50µg	  Cl.	  Notch1	   #2495	   Cell	  Signaling	   1/1000	   50µg	  PDH	   Ab110330	   Abcam	   1/1000	   50µg	  pPDHS293	   NB110-­‐93429	   Novus	  Biologicals	   1/1000	   50µg	  EEA1	   Ab2900	   Abcam	   1/1000	   75µg	  Tubulin	   #2128	   Cell	  Signaling	   1/1000	   Various	  Tubulin	   T0198	   Sigma-­‐Aldrich	   1/1000	   Various	  	  Table 2.4. Antibodies used in western blot experiments.  2.7   Metabolomics Mouse hearts were rapidly exised (<10 s) and freeze-clamped in liquid nitrogen. Frozen hearts were weighed, and homogenized in 375 µL of 50% MeOH per 100 mg tissue weight with the aid of two 5 mm metal balls shaken for 1 min at 30 Hz. Sample temperature was kept below -10°C during tissue homogenization. 375 µL of methanol per 100 mg homogenate weight were then   63 added and the sample was homogenized again for 2 x 1 min. The tube was placed on ice for 30 min and centrifuged at 4°C and 13,000 rpm for 20 min. The supernatant was collected and stored at -20°C for UPLC-FTMS and UPLC-MRM/MS. Carboxylic acids quantification was performed using chemical derivatization with 3-nitrophenylhydrazine (3-NPH) followed by subsequent UPLC-MRM/MS determination according to a protocol as described elsewhere (Han et al., 2013a). Quantitation of glucose, other datable aldoses and reducing sugar phosphates in the mouse heart tissues was performed using chemical derivatization with AEC followed by UPLC-MRM/MS determination using a same protocol as described previously (Han et al., 2013b). Selective quantitation of fructose-1,6-bisP was performed using UPLC-MRM/MS with chemical derivatization. AMP, ADP, and ATP levels were assessed using high-performance liquid chromatography in freeze-clamped hearts. (Ally and Park, 1992). All tissues used in this study were isolated on the same day in the early afternoon.  2.8   Transcriptomics RNA sequencing employed Ion Torrent® technology (Life Technologies, Carlsbad, CA, USA). Specifically, total RNA was quantified using the Qubit™ RNA Assay kit and the Qubit® 2.0 Fluorometer, and its RNA integrity number was measured with the Agilent RNA 6000 Nano kit on the 2100 Bioanalyzer instrument with the 2100 Expert software. Isolation of mRNA was then performed using the Dynabeads® mRNA DIRECT Micro Kit. Whole transcriptome libraries were constructed using the Ion Total RNA-Seq kit on the AB Library Builder™ System. Yield and size distribution of the transcriptome libraries were assess using the Agilent High Sensitivity DNA Kit on the 2100 Bioanalyzer. Templating was constructed using the Ion OneTouch™ 2   64 System using the Ion PI™ Template OT2 200 Kit V3 before sequencing using the Ion PI™ Sequencing 200 Kit V3 on the Ion Proton™ System.  Initial analysis and gene counts were performed on Torrent Suite™ Software (v4.2.1). Downstream analysis was conducted using the Bowtie2/TopHat/CuffLinks/CuffMerge/CuffDiff RNA-Seq analysis pipeline as well as the CumbeRbund software suite in R statistics software (Trapnell et al., 2012).   2.9   Proteomics Hearts were briefly perfused to reduce the number of red blood cells and blood-borne proteins, and then rapidly frozen, prior to proteomic analysis. Frozen hearts were mechanically homogenized and proteins were isolated and treated as previously described (Albu et al., 2015). Isolated protein samples from four individual mice per treatment group were then pooled, quantified, and mixed 1:1 by weight with SILAM heart tissue (Zanivan et al., 2012) that was prepared in the same manner, before final sample processing as described (Albu et al., 2015). Sample digestion and offline fractionation were also conducted as described (Albu et al., 2015) with the exception that endoproteinase LysC was used in place of trypsin for the full digestion. High-pH reverse phase fractionation and mass spectroscopy was conducted as previously described (Albu et al., 2015). Briefly, each sample was fractionated by high-pH reversed phase on an Agilent Zorbax Extend-C18 analytical column (5 µm, 4.6 x 50 mm) into 8 fractions per sample, which were then measured using a Thermo Q-Exactive Hybrid Quadrupole Mass Spectrometer coupled to a Thermo Easy nLC-1000 liquid chromatograph. Each fraction was resolved on a 180-minute gradient and analyzed on the mass spectrometer in positive ion mode, at 70,000 resolution, from 300 to 20,000 m/z. All mass spec data was captured in profile mode.   65 The 10 most intense peaks were selected (underfill ratio of 10%, 2.2 m/z isolation window and a normalized collision energy of 28 with 20% stepping) for fragmentation by HCD, and were excluded thereafter for 30 seconds. Peptides with unassigned charge states, and those with charge 1 were not selected. Peptide identification was carried out using the Andromeda algorithm using the MaxQuant software version 1.5.0.0 from the mouse Uniprot database (July 17th, 2014), largely using the following settings; Endoproteinase LysC cleavage specificity, maximum two missed cleavages, including carbamidomethylcytosine as a fixed modification and protein N-terminal acetylation, methionine oxidation and asparagine and glutamine deamination as variable modifications. Match between runs and depended peptide search were both used at in addition to the default 1% false discovery rate settings. Razor peptides were used for quantification, and MaxQuant re-quantification was enabled. Obvious blood-borne, non-cardiac proteins (e.g. albumin) were ignored. Bioinformatic assoications were visualized using Genemania to determine significantly enriched pathways and potential protein network interactions (Warde-Farley et al., 2010).  2.10   Statistical Analysis Data are expressed as mean ± SEM unless otherwise indicated. Results were considered statistically significant when p < 0.05 using the Student’s t test or two factor mixed design ANOVA with repeated measures and the Bonferroni post-test, where appropriate. All knockout experiments were repeated on at least 3 cRyr2KO mice and at least 3 of their tamoxifen injected control littermates (Ryr2flox/flox ). All induced haploinsufficiency experiments were repeated on at least 3 cRyr2Δ50 mice and at least 3 of their tamoxifen-injected littermate controls (Ryr2flox/wildtype).   66 Chapter 3:  Cardiac RYR2 Controls Heart Rate and Rhythmicity in Adult Mice  3.1   Chapter Summary The molecular mechanisms controlling heart function and rhythmicity are incompletely understood. While it is widely accepted that the type 2 ryanodine receptor (RYR2) is the major Ca2+ release channel in excitation-contraction coupling, the role of these channels in setting a consistent beating rate remains controversial. Gain-of-function RYR2 mutations in humans and genetically engineered mouse models are known to cause Ca2+ leak, arrhythmias and sudden cardiac death. Embryonic stem cell derived cardiomyocytes lacking Ryr2 display slower beating rates, but no supporting in vivo evidence has been presented. The aim of the present study was to test the hypothesis that RYR2 loss-of-function would reduce heart rate and rhythmicity in vivo. We generated inducible, tissue-specific Ryr2 knockout mice with acute ~50% loss of RYR2 protein in the heart, but not in other tissues. Using echocardiography, working heart perfusion, and in vivo ECG telemetry demonstrated that deletion of Ryr2 was sufficient to cause bradycardia and arrhythmia. Our results also show that cardiac Ryr2 knockout mice exhibit some functional and structural hallmarks of heart failure, including sudden cardiac death. These results demonstrate that the RYR2 channel plays an essential role in pacing heart rate. Moreover, we find that RYR2 loss-of-function can lead to fatal arrhythmias typically associated with gain-of-function mutations. Given that RYR2 levels can be reduced in pathological conditions, including heart failure and diabetic cardiomyopathy, we predict that RYR2 loss contributes to disease associated bradycardia, arrhythmia, and sudden death.   67 3.2   Introduction Heart failure is associated with arrhythmias and ends in sudden cardiac death for many patients. The molecular mechanisms linking these pathologies remain to be elicited fully. One candidate is the type 2 ryanodine receptor (RYR2), a large Ca2+ channel in the sarcoplasmic reticulum (SR) membrane. Mutations in the human RYR2 gene cause catecholaminergic polymorphic ventricular tachycardia type 1 (CPVT1) and arrhythmogenic right ventricular dysplasia type 2, rare sudden cardiac death syndromes often manifesting abnormal heart rate (Gomez and Richard, 2004; Priori et al., 2001; Priori and Chen, 2011). Mutations leading to overactive Ca2+ channels and protracted calcium transients have been implicated in most cases (Cerrone et al., 2005; George et al., 2003; Jiang et al., 2005; 2010; Lehnart et al., 2008; Uchinoumi et al., 2010). Gain-of-function ‘knock-in’ mouse models (Chelu et al., 2009; Kannankeril et al., 2006; Kushnir et al., 2010; Uchinoumi et al., 2010; van Oort et al., 2010) confirm non-redundant roles for the RYR2 channel in arrhythmias and sudden cardiac death (Lakatta and DiFrancesco, 2009; Lehnart et al., 2008; 2004b). While these studies suggest that leaky RYR2 channels underlie many arrhythmias, it is controversial whether or not RYR2 mutations alter heart rate. For example, Ryr2R4496C mutant mice develop CPVT1 without a change in basal heart rate (Lakatta and DiFrancesco, 2009). However, mice with point mutations do not permit analyses of other parts of the RYR2 protein or loss-of-function. Recently, potential loss-of-function mutations have been described that are associated with decreased RYR2 open probabilities (Jiang et al., 2007; Thomas et al., 2004). Ryr2 was directly implicated in myocyte beating rate using in vitro differentiated mouse embryonic stem cells (Yang et al., 2002), but the in vivo relevance of these experiments has remained unclear in the absence of inducible Ryr2 knockout mice.   68 The exact mechanisms which set and maintain heart rate remain controversial (Lakatta and DiFrancesco, 2009). One prevalent hypothesis states that heart rate is dictated exclusively by inwardly rectifying channels of the HCN family found in pacemaker cells (Baruscotti et al., 2011). A complementary hypothesis suggests that the electrical events underlying heart rate are governed by two ‘interlocking ionic clocks’ (Lakatta et al., 2010). In this model, heart rate is dependent on an ensemble of interacting ion channels that can be split into two groups, channels of the plasma membrane and channels of the ER/SR, each with their own periods of function and refraction (Lakatta et al., 2010). In this latter model, RYR2 is predicted to be a key regulator of heart rate and rhythmicity in vivo (Honjo et al., 2003a; 2003b; Yang et al., 2002). While vigorous debate remains (Lakatta et al., 2010), there is wide agreement that an in vivo loss-of-function genetic approach is required to test this hypothesis.  Several studies have shown that the levels and/or activity of RYR2 are reduced by up to 50% in disease states such as heart failure and diabetes (Bidasee et al., 2001; Gomez, 1997; Matsui et al., 1995; Naudin et al., 1991). Evidence has also been presented that RYR2 levels diminish with normal aging (Assayag et al., 1998; Tellez et al., 2011). Pathological changes in the expression and/or function of RYR2 channels are usually considered to be downstream of other events in heart disease. Nevertheless, unbiased genome-wide association studies show that RYR2 polymorphisms increase susceptibility to hypertension (Wellcome Trust Case Control Consortium, 2007), meaning that variation in RYR2 expression/function may represent an underlying causal factor in heart disease. Unfortunately, little is known about the in vivo effects of RYR2 loss-of-function. Recently, mice with lifelong Ryr2 haploinsufficiency were found to be resistant to pressure-overload induced cardiac hypertrophy, but also exhibited altered calcium handling, increased cardiomyocyte cell death, and decreased contractility (Zou et al., 2011).   69 Unfortunately, that study of Ryr2+/- mice did not directly examine heart rate or rhythmicity. The lifelong Ryr2 reduction also carries the caveat of possible compensation by other SR calcium channels. In the present study, we tested whether RYR2 has an in vivo role in regulating heart rate and rhythmicity, while confirming the expected role in function. We used inducible, tissue-specific Ryr2 knockout (cRyr2KO) mice, which allowed us to reduce RYR2 in cardiac tissue in vivo in adult animals in a manner that was not associated with compensation from other Ca2+ channels. Our results suggest that RYR2 plays a non-redundant role in the control of heart rate and provide in vivo evidence that RYR2 loss-of-function can lead to arrhythmia.  3.3   Results 3.3.1   Acute reductions in Ryr2 mRNA and protein in conditional knockout mice To create a tissue-specific, inducible model of Ryr2 gene loss-of-function, mice bearing Ryr2flox alleles were crossed with cardiomyocyte-specific, tamoxifen-activated Cre mice (myosin II heavy chain-α promoter) resulting in Ryr2flox/flox:Mer-Cre-Mer mice (cRyr2KO; Fig. 3.1A). Prior to induction, Ryr2 expression was completely normal. However, when treated with the Mer-Cre-Mer agonist tamoxifen, genetic recombination occurred causing Ryr2 gene deletion in adult cardiomyocytes (Fig. 3.1A). Tamoxifen injection for 3 consecutive days resulted in >90% decrease in Ryr2 mRNA, quantified by Taqman RT-qPCR in whole heart, 4 or 10 days after the first tamoxifen dose (Fig. 3.1B). Given that the adult heart contains ~90% cardiomyocytes and ~10% other cell types by volume, these data suggest a virtually complete knockout of the Ryr2 in cardiomyocytes, probably in most regions of these hearts including sinoatrial nodes (Baruscotti et al., 2011). Four days after the initiation of tamoxifen, RYR2 protein was decreased ~50% (Fig.   70 3.1B). RYR2 protein levels declined to ~25% of control values after 10 days (Fig. 3.1B). These observations were further confirmed using immunofluorescence staining, where dramatic changes in the abundance, and the orientation of RYR2 clusters along axial tubules, were evident (Fig. 3.1C). Ryr2 gene knockout was tissue specific (Fig. 3.2A) and we did not observe compensatory increases in other ER calcium release channels (Fig. 3.2B), validating our use of an inducible loss-of-function model to study the role of Ryr2 reduction in heart rate, arrhythmia, and failure.  3.3.2   Sudden cardiac death in cRyr2KO mice Following induction of Ryr2 gene deletion, all cRyr2KO mice developed cardiomyopathy. Four days after the first tamoxifen injection, virtually all cRyr2KO mice were lethargic, but their tamoxifen-injected littermate controls were not. After this time-point, some cRyr2KO mice rapidly reached their humane endpoint whilst others showed a dramatic improvement in health that could last as long as 48 days before eventually reaching their humane endpoint. Interestingly, the final stage had many features of sudden cardiac death. cRyr2KO mice reaching an early humane endpoint showed profound atrial clotting, suggesting the possibility of atrial fibrillation.       71                       Figure 3.1. Acute, cardiac-specific Ryr2 gene ablation in mice. (A) Schemata detailing tamoxifen inducible, cardiomyocyte specific Ryr2 knockout (cRyr2KO) mice and treatment groups. (B) Ryr2 mRNA and RYR2 protein levels in tissues post induction of Ryr2 deletion. Western Blot is representative and plot is quantified against control proteins EEA1 and HSP90. (Ryr2 mRNA: Control n = 6, cRyr2KO n = 6; RYR2 protein: Control n = 3, cRyr2KO n = 3; *p ≤ 0.05). (C) Representative images of disrupted RYR2 protein distribution and re-organization in cryofixed sections of Ryr2 knockout hearts (Control n = 3, cRyr2KO n = 3).   72 Figure 3.2. Conditional Ryr2 knockout is tissue specific and not compensated for by other SR Ca2+ release channels. (A) Ryr2 mRNA levels in heart, brain and skeletal muscle 4 days following induction of Ryr2 deletion (Control n = 6, cRyr2KO n = 6; *p ≤ 0.05). (B) SR Ca2+ release channel mRNA expression 10 days following induction of Ryr2 deletion (Control n = 6, cRyr2KO n = 6; *p ≤ 0.05).  3.3.3   Heart function in cRyr2KO mice To assess cardiac function, we performed M-mode echocardiography on cRyr2KO mice and littermate controls (Fig. 3.3AB). In these experiments, tamoxifen-injected and non-injected Ryr2flox/flox control mice were similar and therefore pooled. The absence of differences in tamoxifen-injected control mice indicated that tamoxifen exposure played no significant role in the profound cardiac phenotype. Acute Ryr2 deletion resulted in a dramatic and rapid >50% decline in cardiac output after just four days (Fig. 3.3C-E). Interestingly, the contractile parameters of cRyr2KO hearts did not appear to decline further over 10 days (Fig. 3.3C-E), and in some tests trended towards normal as part of a compensatory process. To assess cardiac function further in cRyr2KO hearts, we utilized the isolated working heart model (Fig. 3.4A-D). In these experiments, we observed similar decreases in cardiac output, rate pressure product, and hydraulic work demonstrating that these defects were intrinsic to the heart. Thus, Ryr2 is   73 required for normal cardiac function, but there appears to be compensatory mechanisms that transiently ameliorate the detrimental effects of Ryr2 deletion.  Figure 3.3. Conditional Ryr2 knockout mice rapidly lose cardiac function. (A) Timeline of experimental events. (B) Echocardiograms of control and conditional Ryr2 knockout mice. Images are representative (Control n = 7, cRyr2KO n = 5). (C-E) Cardiac output, stroke volume and fractional shortening as measured by echocardiography. Blue lines are control, Red lines are knockout.  (Control n = 7, cRyr2KO n = 5; *p ≤ 0.05).     74  Figure 3.4. Conditional Ryr2 knockout hearts display reduced cardiac function. Measurement of cardiac output (A), peak systolic pressure (B), rate pressure product (C), and hydraulic work (D) in the isolated working heart model (t = 30 min; Control n = 6, cRyr2KO n = 8; *p ≤ 0.05).   3.3.4   Loss of Ryr2 reduces heart rate and results in severe arrhythmias Echocardiography and four-lead ECG were used to assess heart rate in cRyr2KO animals. At times when RYR2 protein levels were ~50% reduced, cRyr2KO mice exhibited a significantly lower rate of heart contraction by echocardiography relative to tamoxifen injected and non-injected littermate controls (Fig. 3.5A). We also observed a concurrent decrease in heart rate, as defined by ECG, suggesting that this drop in heart rate is linked to a decrease in pacemaking at the organ level (Fig. 3.5C). In addition, we observed altered ECG wave patterns in cRyr2KO mice indicating that this decrease in heart rate is genuine sinus bradycardia (Fig. 3.5B). We also observed instances of mice exhibiting a secondary atrioventricular block. This indicates that RYR2 ablation is sufficient to decrease heart rate at both contractile and signaling levels.  cRyr2KO mice were also assessed with the working heart perfusion model to further characterize the link between heart function and heart rate. We consistently observed bradycardia in cRyr2KO hearts having ~50% decrease in RYR2 when compared to tamoxifen treated controls (Fig. 3.5D). A similar decrease in heart rate was observed in all cRyr2KO hearts regardless of their relative cardiomyopathy; hearts in failure showed similar rate decreases to cRyr2KO hearts   75 with relatively normal cardiac function. We observed striking episodes or tachycardic arrhythmia in isolated, working cRyr2KO hearts (Fig. 3.5E). These repetitive arrhythmias were observed in the majority of perfused cRyr2KO hearts throughout the perfusion, although they were more frequently observed early in the perfusion following the stress of isolation and cannulation. Arrhythmias were never observed in tamoxifen-treated control hearts. These working heart experiments suggest that Ryr2 deletion causes an organ-autonomous decrease in heart rate punctuated by brief periods of tachycardic arrhythmia, effects that are independent from systemic factors or neuroregulatory factors in the body. To account for the narrow time frame and to address caveats of echocardiography and working heart perfusion approaches, we studied in vivo heart rate and function in cRyr2KO mice using implantable ECG radio telemetry. Miniaturized two-lead ECG monitors were subcutaneously implanted permitting us to track animal heart rate and activity before and after the induction of gene deletion in cRyr2KO mice. This allowed us to monitor both ECG patterns and heart rate at a very high time resolution in freely moving, unanesthetized animals continuously for extended periods of time. Indeed, we observed a ~20% decrease in average daily heart rate following the induction of cardiac Ryr2 gene deletion when compared to the average daily heart rate of naïve animals (Fig. 3.6A). This decrease in heart rate was not accompanied by any detectable change in average daily activity and was not observed acutely following tamoxifen treatment, suggesting the decrease in heart rate was caused by RYR2 ablation due to gene deletion. During this experiment we also observed periods of repetitive tachycardic arrhythmias at times near humane endpoints (Fig. 3.6B-C). We also observed these arrhythmias following cage changes, a classic stressor of mice, well before humane endpoints and when mice looked healthy by visual inspection. ECG traces showed distinct T-waves and   76 other abnormalities using 2 ECG leads placed on the left pectoral and right ribs, which is state-of-the-art for in vivo mouse telemetry. This study shows that in vivo Ryr2 deletion is sufficient to cause decreased heart rate and episodes of tachycardic arrhythmias in mice.   Figure 3.5. cRyr2 knockout hearts exhibit bradycardia and arrhythmias. (A) Average heart rate of contraction as measured by echocardiography. Measurements were taken at the minimal isoflurane dose necessary to maintain a light plane of anaesthesia. (Control n = 7, cRyr2KO n = 5; *p ≤ 0.05). (B) Four lead ECG of control and cRyr2KO mice taken during echocardiogram. ECG leads were affixed to fore and hind paws. Representative plots (Control n = 7, cRyr2KO n = 5). (C) Quantification of heart rate as measured by ECG during echocardiography experiments (Control n = 6, cRyr2KO n = 4; *p ≤ 0.05). (D) Average heart rate of isolated, perfused working hearts (Control n = 5, cRyr2KO n = 5; *p ≤ 0.05). (E) Examples of heart rate traces from isolated, perfused working hearts. Heart rate calculated from changes in cardiac output measured by flow meters. All cRyr2KO hearts exhibited dramatic periodic arrhythmias that were never observed in control hearts (Control n = 5, cRyr2KO n = 5).       77                    Figure 3.6. cRyr2 knockout mice exhibit bradycardia and arrhythmias in vivo. (A) Average heart rate collected from freely moving, unanaesthetized cRyr2KO mice using implantable ECG telemetry (n = 4, initially). White points denote pre-knockout timepoints, red points denote the days of tamoxifen injections, and black points denote days following gene knockout. Red X’s indicate days when mice reached humane endpoints. Heart rate data is expressed as a percentage of the average heart rate of pre-knockout days, which is depicted as a dashed blue line (*p ≤ 0.05 when compared to average naive heart rate). (B) Representative ECG traces from a single cRyr2KO knockout mouse prior to gene knockout (blue) and during a characteristic arrhythmic event (red) observed 20 days following gene knockout (n = 4). (C) Enlarged regions of ECG traces seen in panel B. Green roman numerals and underlines designate enlarged regions.    78 3.4   Discussion In this study we used inducible, tissue-specific gene ablation to examine the in vivo functions of Ryr2 in the adult heart. Inducible Ryr2 deletion circumvented chronic gene compensation and the embryonic lethality of the global Ryr2-/- mice. This is the first reported tissue-specific deletion of any RYR isoform in any tissue. This model permitted the first in vivo and ex vivo analysis of specific cardiac Ryr2 loss-of-function in mice. Our in vivo data shed light on controversial roles for this gene in heart rate and rhythmicity, while confirming expected cardiac dysfunction following Ryr2 deletion. Specifically, we found that Ryr2 is required for setting normal heart rate, and that deletion of this gene causes general bradycardia. We further found that Ryr2 is required to maintain normal heart rhythm, and that Ryr2 deletion causes intermittent tachycardic arrhythmias. These results indicate that Ryr2 has a non-redundant role in regulating heart rate. We also confirmed that a reduction in RYR2 protein is sufficient to induce aspects of heart failure in mice. These effects were observed following Ryr2 deletion and in the absence of other stressors. Although tamoxifen, the drug used to initiate Mer-Cre-Mer mediated gene deletion, is known to have mild cardiac effects, our use of tamoxifen-injected control mice excludes the possibility that tamoxifen plays any significant role in the observed phenomena.  Our data support the hypothesis that RYR2 has a non-redundant role in controlling heart rate as we observed a consistent decrease in global heart rate following induction of gene deletion both in vivo and ex vivo. Our results are consistent with previous in vitro work. For example, ryanodine, a pharmacological modulator of RYR2 opening probability, is sufficient to alter the contraction rate of atrial tissue cultures (Honjo et al., 2003b). Similarly, model cardiac cells derived from mouse embryonic stem cells lacking Ryr2 show a reduced rate of spontaneous contraction (Yang et al., 2002). Our results provide dramatic evidence that RYR2 does indeed   79 play this role on an organ level in adult cardiac tissues, in vivo and ex vivo. As such, our results support the “calcium clock” hypothesis, which predicts that the periodicity of Ca2+ transients at the level of the SR/ER, a process governed in part by RYR2, is critical to the electrochemical signaling underlying heart rate (Lakatta et al., 2010). It is worth noting that our results do not exclude parallel or sequential involvement of other cardiac ion channels (e.g. HCN) in the regulation of heart rate. Our data suggests that in vivo deletion of Ryr2 may have altered the rate at which pacemaking cells signaled the initiation of heart contraction or in the manner cardiac action potentials are conducted. However, we cannot yet conclusively say whether the effects are dependent on nodal cells, conduction tissues, or primarily atrial and ventricular cardiomyocytes, or some combination. The model system that we employed relies on the α-MHC promoter to drive Cre recombinase expression and gene deletion, which is abundantly expressed in both atrial and ventricular cells in young adult mouse hearts. Unfortunately, it is not technically feasible for us to quantitative assess RYR2 protein levels specifically in isolated sinoatrial nodes of cRyr2KO mice and littermate controls by immunoblot. However, a recent study using the same C57Bl6-mhy6-Mer-Cre-Mer transgenic mouse strain to drive gene knockout broadly in the heart confirmed robust cre-lox recombination in the sinoatrial node (Baruscotti et al., 2011). Given that we used the identical Cre-deleter mouse model, it is reasonable to expect that a similar degree of Ryr2 gene deletion occurred in the sinoatrial nodes of our mice.  Our results also reveal that decreased RYR2 activity can cause tachycardic arrhythmias. RYR2 has a well-documented role in the genesis of arrhythmias as mutations in the human RYR2 gene underlie tachycardic arrythmogenic conditions such as CPVT and ARVD (George et al., 2007a). The vast majority of mutations studied are predicted to increase RYR2 open probabilities   80 by increasing the channels sensitivity to cytosolic or SR luminal Ca2+ levels (George et al., 2007a; Jiang et al., 2005; Tung et al., 2010). Thus, these Ryr2 mutants are considered “leaky” as they produce prolonged or inappropriate calcium transients (George et al., 2003; Jiang et al., 2004; Lehnart et al., 2004b; Thomas et al., 2004). The prevailing model for how Ryr2 mutations cause CPVT suggests that aberrant, increased RYR2 activity during diastole drives NCX currents in reverse (Na+ influx, Ca2+ efflux) potentiating delayed after-depolarizations which can initiate rapid, arrhythmic heart contractions in a feed forward manner (Pogwizd, 2004). However, this model cannot explain our observation of tachycardic arrhythmias following Ryr2 deletion and reduced RYR2 activity. Therefore, an alternate mechanism must underlie observed ventricular fibrillation in cRyr2KO mice following gene deletion.  Previous studies conducted on L433P and A4860G Ryr2 mutants, known to cause either catecholaminergic idiopathic ventricular fibrillation or sudden cardiac death respectively, found decreased RYR2 open probability and insensitivity to SR luminal Ca2+ levels (Jiang et al., 2007; Thomas et al., 2004). The authors of those papers suggested that these mutations may potentiate arrhythmias via an alternans-dependent mechanism (Jiang et al., 2007; Thomas et al., 2004). Alternans, in this case, refer to alternating strong and weak cellular calcium transients which are associated with heart failure and ventricular fibrillation (Hüser et al., 2000; Picht et al., 2006; Surawicz and Fisch, 1992). Although a variety of stimuli can cause cellular alternans: such as metabolic alterations, acidosis, and certain electrophysiological stimulations, it is thought that virtually all alternans are caused by alterations in RYR2 behavior (Hüser et al., 2000; Picht et al., 2006; Surawicz and Fisch, 1992). One study used tetracaine, an inhibitor of RYR2 action, to induce alternans by decreasing RYR2 activity which functionally uncoupled groups of RYR2 from one another leading to heterogeneous subcellular Ca2+ waves which alternated spatially and   81 temporally (Diaz et al., 2002). It is possible that this phenomenon may lead to prolonged or inappropriate calcium transients which could potentially lead to delayed after-depolarizations and arrhythmias. While our research does not describe the precise mechanism by which Ryr2 deletion causes arrhythmias, our results are consistent with the model of alternans-associated ventricular fibrillation caused by decreased RYR2 activity (Jiang et al., 2007; Thomas et al., 2004). Overall though, our research in combination with other work suggests that appropriate RYR2 function, that is neither elevated nor decreased, is essential for preventing arrhythmias. It is thought that up to 50% of heart failure associated deaths are caused, in an acute sense, by arrhythmias (Kannel et al., 1988). It is also thought that these arrhythmias are caused by cardiac alternans (Surawicz and Fisch, 1992), which are frequently observed during heart failure (Euler, 1999; Kodama et al., 2001), and are known to be a pathology of RYR2 dysfunction (Diaz et al., 2002; Zou et al., 2011). In addition, decreased RYR2 levels or reduced RYR2 activity have been observed in several models of heart failure, whether induced by pressure overload (Matsui et al., 1995; Naudin et al., 1991), chronic hypertension (Gomez, 1997), or diabetes (Bidasee et al., 2001). The degree of RYR2 dysfunction correlates with the degree of cardiac dysfunction (Belevych et al., 2011). Whether such decreases in RYR2 function are sufficient to cause heart failure, or are merely consequences of pathological dysfunction remains a somewhat controversial idea because RYR2 loss is not observed in all studies. However, our results provide compelling evidence that decreased RYR2 function can contribute to heart failure in vivo and that Ryr2 deletion alone is sufficient to recapitulate functional characteristics of heart failure: reduced contractility, decreased cardiac output, and ventricular fibrillation. Our results, along with other work (Diaz et al., 2002; Jiang et al., 2007; Thomas et al., 2004), also suggest that the decreased RYR2 function observed in heart failure may be responsible for ventricular fibrillation   82 caused by subcellular Ca2+ alternans. Collectively, our research suggests that RYR2 may be a key player in cardiac dysfunction during heart failure.    83 Chapter 4:  Cardiomyocyte ATP Production, Metabolic Flexibility, and Survival Require Calcium Flux Through Cardiac Ryanodine Receptors In Vivo  4.1   Chapter Summary Ca2+ fluxes between adjacent organelles are thought to control many cellular processes, including metabolism and cell survival. In vitro evidence has been presented that constitutive Ca2+ flux from intracellular stores into mitochondria is required for basal cellular metabolism, but these observations have not been tested in vivo. Here, we report that controlled in vivo depletion of cardiac RYR2, using a conditional gene knockout strategy (cRyr2KO mice), is sufficient to reduce mitochondrial Ca2+ and oxidative metabolism, and to establish a pseudo-hypoxic state with increased autophagy. Dramatic metabolic reprogramming was evident at the transcriptional level via Sirt1/Foxo1/Pgc1α, Atf3, and Klf15 gene networks. Ryr2 loss also induced a non-apoptotic form of programmed cell death associated with increased calpain-10, but not caspase-3 activation or ER-stress. Remarkably, cRyr2KO mice rapidly exhibited many of the structural, metabolic and molecular characteristics of heart failure at a time when RYR2 protein was reduced 50%, a similar degree to that which has been reported in heart failure. RYR2-mediated Ca2+ fluxes are therefore proximal controllers of mitochondrial Ca2+, ATP levels, and a cascade of transcription factors controlling metabolism and survival.       84 4.2   Introduction Intracellular Ca2+ fluxes regulate an enormous number of processes with the specificity of responses often being ensured by spatial limitation of Ca2+ ions and proximity to targets. Frequently, this takes the form of Ca2+ signals within the nanoscale space between adjacent organelles (Johnson et al., 2012). For example, in vitro studies have shown that Ca2+ flux through channels such as ryanodine receptors and IP3-receptors mediate privileged communication between endoplasmic reticulum/sarcoplasmic reticulum (ER/SR) and mitochondria and that this paces cellular metabolism by stimulating oxidative ATP production via interaction with TCA cycle enzymes (Denton and McCormack, 1990; Giacomello et al., 2010; Liu and O'Rourke, 2008).  Elegant work from Foskett and colleagues recently demonstrated that knocking out IP3Rs in DT40 cells causes an energy-deficient state and mTOR-independent autophagy (Cárdenas et al., 2010). This concept has not been extended to other cell types or to the in vivo situation.  Ryanodine receptor Ca2+ channels (e.g. RYR2) have been observed in close proximity to mitochondria, and beat-to-beat calcium transients have been observed in cardiomyocyte mitochondria (Kettlewell et al., 2009; Lukyanenko et al., 2007; Salnikov et al., 2009). Furthermore, we and others have demonstrated in vitro that RYR2-mediated Ca2+ flux supports ATP levels in other cell types (Dror et al., 2008; Glancy and Balaban, 2012; Johnson, 2004; Tsuboi et al., 2003). In pancreatic β-cells, blocking RYR2 also induces calpain-10-dependent, caspase-3-independent, programmed cell death that is associated with presenilin1-dependent upregulation of hypoxia-inducible factor 1β (Dror et al., 2008; Johnson, 2004). To date, analysis of cellular energetics and survival in the context of reduced ER/SR Ca2+ channel expression has not been extended to an in   85 vivo system. Ideally, the hypothesis that intracellular Ca2+ release is required to sustain energy metabolism in vivo would be tested using tissue-specific, inducible deletion of ER/SR Ca2+ channels. The heart is an ideal model to test this hypothesis because it is a tissue with high metabolic demands. Notably, a single intracellular Ca2+ release channel type, RYR2, is far in excess of other analogous Ca2+ channels, meaning that ER/SR Ca2+ release may be significantly reduced with a single genetic mutation. Examination of cardiomyocytes with specific Ryr2 gene loss-of-function is also of interest because RYR2 protein levels are known to decline as much as 50% with age, in parallel with an increased risk for heart failure (Assayag et al., 1998; Kandilci et al., 2011). RYR2 expression, structural organization and function are reduced in heart failure, cardiac hypertrophy and ischemic cardiomyopathy (Bidasee et al., 2001; Brillantes et al., 1992; Matsui et al., 1995; Naudin et al., 1991). It is unclear whether RYR2 loss is sufficient to cause any of the features of heart failure, including contractile dysfunction, hypertrophy, fibrosis, inflammation, transcriptional reprogramming, an energy-starved state of metabolic inflexibility, and cardiomyocyte death (Crossman et al., 2011; Ferrari et al., 2009; Stanley, 2005).  In the present study, we demonstrate that acute in vivo deletion of the primary ER/SR Ca2+ release channel reduced ATP, blunted oxidative mitochondrial metabolism, and led to inflexibility of energy substrate utilization. We provide evidence that this reduced energy state associated with loss of RYR2-mediated Ca2+ flux, activates hypoxia-inducible factors and switches off transcriptional control hubs that govern a wide range of key metabolic pathways. We further observed atypical Calpain-10-dependent cell death and cardiac hypertrophy in mice following Ryr2 knockout. Acute in vivo deletion of cardiomyocyte Ryr2 reproduces many   86 hallmarks of heart failure, suggesting a causal, upstream role for this channel in the pathogenesis of this disease.  4.3   Results 4.3.1   Ryr2 deletion reduces Ca2+ flux in cardiac myocytes  cRyr2KO mice showed normal Ryr2 expression until they were injected with tamoxifen, a synthetic agonist of the modified estrogen receptor:Cre fusion protein (Mer-Cre-Mer) under the control of a αMhy6 promotor, which allowed temporally controlled targeted gene deletion of Ryr2 in cardiomyocytes. Tamoxifen injected littermate control mice (Ryr2flox:flox) were used throughout the study, thus any differences between cRyr2KO and control mice cannot be attributed to tamoxifen. In our hands, this model consistently displays a >90% decrease in whole heart Ryr2 mRNA after only 3 days of tamoxifen injections (Fig. 4.1A) (Bround et al., 2012). This leads to a time-dependent reduction in RYR2 protein levels to 51 ± 21% at 4 days post tamoxifen and 24 ± 22% at 10 days post tamoxifen in cRyr2KO mouse hearts (Bround et al., 2012). No effects were seen in the expression of Ryr1, Ryr3, or any of the IP3 receptor Ca2+ release channels (Fig. 4.1A). As expected for cells lacking their major Ca2+ release channel and without compensatory upregulation of other SR channels, steady state cytosolic Ca2+ was reduced in isolated, acutely cultured, non-contracting cRyr2KO cardiomyocytes (Fig. 4.1B). We predicted that the loss of RYR2 channels juxtaposed with mitochondria would result in lower average Ca2+ levels in mitochondria. Indeed, we observed a significant decrease in steady state mitochondrial Ca2+ in isolated cRyr2KO cardiomyocytes (Fig. 4.1C). This result was not dependent on the normalization of the Rhod2 signal to Mitotracker fluorescence (used to control for mitochondrial number). Together, these studies suggest that loss of RYR2 in cardiomyocytes   87 reduces Ca2+ flux from the SR into the mitochondria, via the cytosol.   4.3.2   RYR2 supports ATP production in vivo In vitro studies have shown that Ca2+ flux from ER/SR stimulates metabolism in adjacent mitochondria (Giacomello et al., 2010; Glancy and Balaban, 2012; Liu and O'Rourke, 2008). We used cRyr2KO hearts to test the hypothesis that SR-to-mitochondria Ca2+ fluxes are required for ATP production and oxidative metabolism in vivo. Indeed, ATP levels were decreased in flash-frozen cRyr2KO hearts (Fig. 4.2A). Consistent with this model, metabolic profiling of cRyr2KO working hearts revealed significant reductions in glucose, palmitate, and lactate oxidation (Fig. 4.2B). These effects were specific to mitochondrial metabolism, as we did not observe a reduction in the rate of glycolysis (Fig. 4.2C). In fact, glycolysis was increased in cRyr2KO working hearts, suggesting a compensatory mechanism and a switch towards a heart failure-like phenotype (Stanley, 2005). These observations were made in hearts with identically oxygenated perfusate and with similar amounts of coronary artery perfusion indicating that the cardiomyocytes had similar access to metabolic substrates (Fig. 4.2D). Together, these data provide evidence that Ca2+ flux through RYR2 channels is essential for basal mitochondrial metabolism in the heart.      88  Figure 4.1. Acute, cardiac-specific Ryr2 gene ablation without compensation from related Ca2+ channels. (A) SR/ER Ca2+ release channel mRNA levels in heart tissue 10 days after tamoxifen injection. Data is normalized to cyclophilin gene levels (n = 6). White bars = Control mice, Black bars = cRyr2KO mice throughout. (B) Reduced basal cytosolic Ca2+ levels in isolated, acutely cultured, un-stimulated cardiomyocytes from cRyr2KO mice. Data are from at least 3 independent cell preparations (Control cells n = 124; cRyr2KO cells n = 210). (C) Reduced basal mitochondrial luminal Ca2+ levels in isolated, un-stimulated cardiomyocytes from cRyr2KO mice. Data are from 4 independent cell preparations (Control cells n = 170; cRyr2KO cells n = 180; * denotes p ≤ 0.05). White scale bars are the equivalent 20 µm. * denotes p ≤ 0.05.    89           Figure 4.2. Loss of RYR2-mediated Ca2+ flux reduces mitochondrial metabolism and steady-state ATP levels. (A) ATP levels of freshly excised and freeze-clamped hearts (n = 9). White bars = Control, Black bars = cRyr2KO. (B) Oxidation rates of glucose, palmitate, and lactate by isolated, perfused working hearts (Control n = 3; cRyr2KO n = 4). (C) Rates of glycolysis during working heart preparation (Control n = 3; cRyr2KO n = 4). (D) Coronary arterial flow during the working heart perfusions when metabolic assessments were made (Control n = 6; cRyr2KO n = 8). All data collected 4 days post first tamoxifen injection. * denotes p ≤ 0.05.   4.3.3   Ryr2 deletion leads to a hypoxic/ischemic state Our previous in vitro work on pancreatic β-cells demonstrated that the chemical inhibitor of RYRs, ryanodine, caused a state of ATP depletion and a Presenilin-1-dependent induction of Hypoxia-inducible factor (Dror et al., 2008). The Hypoxia-inducible factor system is a master sensor of energy use and may therefore coordinate the response to energy deprivation. Here, we also observed an increase in the expression of presenilin-1 and presenilin-2, as well as HIF1α and HIF1β (Fig. 4.3A,B). A dramatic loss of uncoupling protein gene expression suggested that cRyr2KO cardiomyocytes were reprogrammed to conserve and salvage energy (Fig. 4.3C).   90 Similar to the in vitro observations of others (Cárdenas et al., 2010), we observed increased autophagy under conditions of nutrient availability, based on increased levels of LC3-II (Fig. 4.3D). Together, these observations suggest that cardiomyocytes display complex signs of energy stress and compensation when ATP levels are reduced following RYR2 reduction.     Figure 4.3. Acute Ryr2 ablation causes a hypoxia-like cellular state. (A) Presenilin-1 (22 and 55 kDa) and presenilin-2 (23 and 54 kDa) protein levels. White bars = Control, Black bars = cRyr2KO throughout.  Individual blot lanes are biological replicates throughout (n = 3). (B) HIF1α (120 kDa) and HIF1β (95 kDa) protein levels (HIF1α n = 3; HIF1β n = 7). (C) Uncoupling protein mRNA (n = 3). (D) Autophagy marker LC3-I (18 kDa) and LC3-II (16 kDa) (n = 3). Protein quantification plots represent relative expression normalized to tubulin (50 kDa). All data collected 4 days post first tamoxifen injection. * denotes p ≤ 0.05.      91 4.3.4   Reprogramming of metabolism in cRyr2KO hearts Given the severity of the metabolic changes, we next tested whether core transcriptional pathways had been affected by the reduction in RYR2-mediated Ca2+ fluxes. We focused our efforts on so-called ‘master regulator’ pathways that modulate the expression of numerous metabolic genes in heart and other tissues. For example, we observed downregulation of Sirt1 and Foxo1 (Fig. 4.4A), genes that coordinate cardiac stress responses and metabolism (Buteau et al., 2007; Nemoto et al., 2004; Planavila et al., 2011). Heart failure is associated with metabolic inflexibility and a switch away from fatty acid metabolism (Lionetti et al., 2011). Ca2+ signals can act via calcineurin-dependent activation of the Ppargc1a (Pgc1α)/Pparα system to sustain fatty acid oxidation (Schaeffer et al., 2004; Son et al., 2010). In hearts with reduced RYR2, we observed a striking downregulation of Ppargc1a and Pparα, as well as Pparγ (Fig. 4.4B). Phosphoenolpyruvate carboxykinase 1 (Pck1), a target of Pgc1α, was virtually eliminated from cRyr2KO hearts (Fig. 4.4G). The Pparg target gene Cebpa was also robustly inhibited (Fig. 4.4C). Carbohydrate responsive element-binding protein (Chrebp), a master regulator of complex lipid metabolism, was dramatically decreased in cRyr2KO hearts (Fig. 4.4C). Accordingly, expression of ATP-citrate lyase (Acly) and acetyl-CoA carboxylase β (Acacb) were decreased significantly; expression of stearoyl-Coenzyme A desaturase 1 (Scd1) was virtually eliminated (Fig. 4.4D,E). The reduction in lipogenic enzymes was associated with downregulation of fat transporters (Fig. 4.4F). At the same time, we observed a dramatic decrease in expression of genes encoding hormone sensitive lipase (Hsl), adiponutrin (Adpn), and adipose triglyceride lipase (Atgl) (Fig. 4.4H). Atgl is essential for degrading lipid droplets and mice lacking this gene have fatty, hypertrophied, failing hearts (Haemmerle et al., 2006).   92 Interesting, we noted reduced expression of the genes encoding the GLUT4 glucose transporter and phosphoenolpyruvate carboxykinase 1 (Fig. 4.4G,I). Together, these data suggest that the use of lipids and other substrates by the cRyr2KO heart may be abrogated and provide novel insight into the link between Ca2+ and lipid metabolism.              Figure 4.4. Conditional Ryr2 knockout results in metabolic reprogramming at the transcriptional level. (A-J) Expression of key metabolic gene mRNA in whole heart was measured 4 days post-tamoxifen using Taqman RT-qPCR. We measured mRNA expression levels of sirtuin 1 (SIRT1) (A), forkhead box O1 (FOXO1) (A), peroxisome proliferator-activated receptor gamma, coactivator 1 alpha (Ppargc1a) (B), peroxisome proliferator-activated receptor gamma (Pparg) (B), peroxisome proliferator activated receptor alpha (Ppara) (B), CCAAT/enhancer binding protein (C/EBP), alpha (Cebpa) (C), sterol regulatory element binding transcription factor 1 (SREBP1c) (C), Carbohydrate response element-binding protein (ChREBP) (C), ATP citrate lyase (ACLY) (D), acetyl-CoA carboxylase beta (ACACB) (D), stearoyl-Coenzyme A desaturase 1 (SCD1) (E), fatty acid binding protein 4, adipocyte (Fabp4) (F), CD36 molecule (CD36) (F), phosphoenolpyruvate carboxykinase 1, cytosolic (Pck1) (G), hormone sensitive lipase (HSL) (H), adiponutrin (ADPN) (H), adipose triglyceride lipase (ATGL) (H), fatty acid synthase (FASN) (H),  and solute carrier family 2 (facilitated glucose transporter), member 4 (GLUT4) (I). White bars = Control, Black bars = cRyr2KO throughout (n = 3). All data were collected 4 days post first tamoxifen injection. * denotes p ≤ 0.05.    93 4.3.5   Ryr2 deletion induces programmed cell death Heart failure is associated with an increase in cardiomyocyte death, which can be apoptotic, non-apoptotic and/or associated with ER-stress, depending on the model (Ferrari et al., 2009). cRyr2KO hearts contained numerous TUNEL-positive cells, which were virtually absent in controls (Fig. 4.5A,B). This cell death was not caused by ER-stress, since we observed a decrease in CHOP and other ER-stress markers (Fig. 4.5C,D). In some cell types, RYR2 hyperactivity contributes to classical caspase-3-dependent apoptosis (Luciani et al., 2008). On the other hand, we previously discovered that inhibiting RYR2-mediated Ca2+ flux with ryanodine caused caspase-3-independent programmed cell death (Johnson, 2004). cRyr2KO hearts did not exhibit a significant increase in caspase-3 cleavage (Fig. 4.5E), confirming our hypothesis that RYR2 suppresses an atypical form of non-apoptotic cell death. We observed a reduction in the expression of the pro-survival gene Bcl2, perhaps further contributing to cell death (Fig. 4.5F). We have previously shown that calpain-10 mRNA and small molecular weight isoforms of calpain-10 are upregulated in cells with depressed energy state resulting from ryanodine treatment (Dror et al., 2008; Johnson, 2004). Two protein isoforms of calpain-10 were significantly increased in heart tissue from cRyr2KO mice (Fig. 4.5G). These bands likely represent calpain-10 protein isoforms and cleavage products, as they are not observed in tissue from calpain-10 knockout mice (Johnson, 2004). No effects were observed on the levels of calpain-1 or calpain-2 protein in cRyr2KO hearts. Thus, loss of Ryr2 function activates a conserved form of calpain-10-associated programmed cell death in multiple tissues, including the heart.      94                                      Figure 4.5. Acute Ryr2 ablation causes calpain-10 dependent programmed cell death. (A) Representative imagines of TUNEL staining (arrows) of fixed heart sections (Scale Bar=100µm). (B) Quantification of TUNEL positive nuclei (Control n = 4, cRyr2KO n = 5). White bars = Control, Black bars = cRyr2KO throughout. (C) ER stress gene expression (n = 3). (D) CHOP (31 kDa) protein expression (n=7). (E) Cleaved caspase-3 (19 kDa) protein levels. + denotes a positive cleaved caspase-3 control sample (thapsigargin-treated MIN6 cells). Individual blot lanes are biological replicates throughout (n = 3). (F) Cell death effecter gene expression 4 days post tamoxifen injection (n = 3). (G) Calpain-10 protein levels and expression patterns. Calpain-1 (80 kDa) and Calpain-2 (80 kDa) levels are also displayed. White bars = Control, Red bars = cRyr2KO.  (H) Insulin receptor (Insr) and Ptpn1 gene expression levels (n = 3). (I) Adiponectin and adiponectin receptor mRNA expression. (n = 3) (J) Leptin receptor mRNA (n = 3). (K) Expression and/or phosphorylation states of Insulin receptor (190 kDa, 95 kDa), Akt (60 kDa), and Erk (42 and 44 kDa) (n = 3;). All analyses carried out 4 days post-tamoxifen treatment. Protein quantification plots represent relative expression normalized to either total protein or to tubulin (50 kDa). TUNEL data were collected 10 days following tamoxifen treatment; all other data collected 4 days following first tamoxifen injection. * denotes p ≤ 0.05.   95 4.3.6   Loss of cardioprotective pathways in cRyr2KO hearts Cell death can be accelerated when growth factor signaling pathways are down-regulated, and hypoxia-like conditions have been linked to insulin resistance (Regazzetti et al., 2009). cRyr2KO hearts had a significant reduction in the mRNA and protein levels of the insulin receptor (Fig. 4.5H,K). A trend towards increased Ptpn1 expression further suggested a suppression of insulin signaling (Fig. 4.5H), and this was supported by a significant reduction in ERK activity and pro-survival AKT phosphorylation at threonine 308 (Fig. 4.5K). Cardioprotective adiponectin and leptin signaling systems were downregulated (Fig. 4.5I,J). These data suggest that survival pathways are reduced following Ryr2 reduction.   4.3.7   Ryr2 deletion causes cardiac hypertrophy and fibrosis cRyr2KO mice represented an ideal tool to test the hypothesis that acute cardiomyocyte stress and dysfunction is sufficient to induce cardiac hypertrophy. We observed an increase in gross heart volume and mass (Fig. 4.6A) and heart fibrosis in cRyr2KO mice (Fig. 4.6B), already evident 10 days after tamoxifen injection. Many chronic models of cardiac hypertrophy result in a re-expression of fetal genes, including cardioprotective growth factors. In our acute model, brain natriuretic peptide transcription was significantly increased and adult β-myosin heavy chain gene (Mhy6) was attenuated (Fig. 4.6C-D), but we did not find a simultaneous upregulation of the fetal β-myosin heavy chain gene (Mhy7; Fig. 4.6D). Interestingly, we observed a reduction of cleaved Notch1 (Fig. 4.6E), which plays a key role in maintaining cardiac progenitors (Croquelois et al., 2008). Heart failure can be associated with an increase in cardiac inflammation and the local release of pro-inflammatory cytokines (Rohini et al., 2010). We observed a dramatic 20-fold increase in interleukin-6 transcription in cRyr2KO hearts (Fig.   96 4.6F). Ryr2 may therefore be upstream of some, but not all, aspects of cardiac inflammation.  Next, we examined pathways that could mediate translation of the effects of stress into the hypertrophic pathology. Atf3, a stress gene activated by ischemia/hypoxia (Huang et al., 2008; Igwe et al., 2009), was increased ~2.5-fold in cRyr2KO hearts (Fig. 4.6G). Cardiac Atf3 overexpression is sufficient to induce heart failure, hypertrophy and myocyte death (Okamoto et al., 2001). We also observed a dramatic loss of several genes negatively regulated by Atf3, including Glut4 (Fig. 4.4I). cRyr2KO also resulted in a near complete loss of Klf15 (Fig. 4.6H) which, based on knockout studies, is sufficient to induce heart failure and fibrosis (Fisch et al., 2007; Koh et al., 2010). Klf15 is a positive regulator of Ucp1 and Pparg (Yamamoto et al., 2010), genes that were decreased in cRyr2KO hearts. Thus, acute Ryr2 knockout and the concomitant energy deficient state have profound effects on transcriptional hubs, controlled, at least in part, by hypoxia-inducible factors, Klf15, Atf3, and Foxo1/Sirt1. Throughout this study, we assessed the survival of cRyr2KO mice. Despite molecular and/or physical compensation, cRyr2KO mice invariably met their humane endpoint after a period that could range from days to weeks, as we have shown previously (Bround et al., 2012). Thus, the cardiac expression of the Ryr2 gene is essential for survival in adult mice.         97   Figure 4.6. Cardiac hypertrophy and fibrosis in cRyr2KO hearts. (A) Heart size, heart wet weight relative to body mass, and total body mass 10 days following tamoxifen treatment (Control n = 6, cRyr2KO n = 8; *p ≤ 0.05). Picture is representative. (B) Masson's Trichrome staining of cardiac tissue sections collected ten days following tamoxifen treatment. Lower panels are higher magnification images of top panels. Images are representative (Control n = 4, cRyr2KO n = 5; Scale bar = 100 µm). (C,D) Anp, Bnp, Mhy6 and Mhy7 gene expression 4 and 10 days after tamoxifen treatment. (n = 3) (E) Cleaved Notch1 (110 kDa) protein at 4-days post-tamoxifen. Individual blot lanes are biological replicates throughout (n = 3). (F) Inflammatory gene expression 4 days post-tamoxifen (n = 3). (G-H) Gene expression levels of ATF3 and KLF15 assessed 4 days post-tamoxifen (n = 3). Protein quantification plots represent relative expression normalized to either total protein or to tubulin (50 kDa). * denotes p ≤ 0.05.      98 4.4   Discussion The purpose of this study was to test the hypothesis that Ca2+ flux from intracellular stores, in this case through cardiomyocyte SR RYR2 channels, is required to maintain cellular energy homeostasis and survival in vivo. We used tissue-specific, inducible gene ablation to examine the in vivo functions of Ryr2 in the adult mouse heart. This model circumvents the embryonic lethality of Ryr2-/- mice (Takeshima et al., 1998), as well as the possible long-term compensation from related Ca2+ channels reported in Ryr2+/- mice (Zou et al., 2011). Our in vivo mouse studies support the concept, previously proposed after in vitro studies (Cárdenas et al., 2010), that intracellular Ca2+ release paces mitochondrial metabolism and protects cells from atypical programmed cell death caused by energy depletion. Our data extend this concept to include the Ca2+-dependent control of key transcription factor networks that modulate metabolic substrate utilization. Our results also demonstrate that partial loss of cardiomyocyte RYR2 protein is sufficient to recapitulate many of the characteristics of human heart failure, providing new mechanistic insight into this devastating condition. In the physiological state, the cardiomyocyte is always working. Our analysis of metabolism and survival were conducted in vivo, and in ex vivo working heart models. The single-cell Ca2+ measurements were the only experiments in this study not conducted in working cardiomyocytes. Nevertheless, these measurements provide some interesting information and suggest a role for RYR2 channels in cardiomyocyte Ca2+ homeostasis. Taken at face value, these data suggest that basal Ca2+ flux through RYR2 channels normally supports a significant component of resting Ca2+ levels in both the cytosolic and mitochondrial compartments of isolated cardiomyocytes. Indeed, microscopic and sub-microscopic RYR2-mediated Ca2+ release events have previously been implicated in the diastolic SR Ca2+ leak into the cytosol (Santiago et   99 al., 2010), and sub-threshold RYR2 release events are known to increase Ca2+ in nearby mitochondria (Pacher et al., 2002). Future studies will be required to assess Ca2+ in these compartments in beating cardiomyocytes, ideally within the intact heart. One of the key findings of our study is that cRyr2KO mouse hearts had decreased ATP levels, as well as lower substrate utilization when compared to control hearts. This can be attributed to defects in the utilization of metabolic substrates via oxidative ATP generation. The Ca2+ dependence of the TCA cycle was proposed many years ago based on the finding that Ca2+ in the matrix of isolated mitochondria stimulates several enzymes in the pathway, including pyruvate dehydrogenase, which controls entry of glucose carbon into the TCA cycle (Denton and McCormack, 1985; 1990; Wan et al., 1989). Similarly, emerging evidence suggests that the F1F0 ATP synthase is also directly regulated by mitochondrial Ca2+ such that the maximal rate of oxidative ATP production at given mitochondrial membrane potential is dependent on Ca2+ concentration (Glancy and Balaban, 2012). As such, the rate and efficacy of many energy-producing substrates via the TCA cycle could be affected by mitochondrial Ca2+ dynamics. Collectively, these enzymes are thought to rely on Ca2+ signals from across nanoscale spaces between ER/SR and mitochondria. Local microdomains of concentrated Ca2+ are required to access the mitochondria and the duration of the mitochondrial Ca2+ transients is limited by rapid export mechanisms (Giacomello et al., 2010; Kettlewell et al., 2009; Liu and O'Rourke, 2008). Our results suggest that ablation of RYR2 in vivo disrupts this signaling pathway, leading to a decrease in TCA cycle enzymatic activity and consequently a diminished ability to sustain cellular ATP concentrations. Inability to employ the TCA cycle likely promotes the metabolic inflexibility observed in heart failure, as the utilization of fat as fuel requires oxidative   100 metabolism (Stanley and Chandler, 2002). Indeed, decreased RYR2 expression or function is associated with several models of cardiomyopathy (Bidasee et al., 2001; Brillantes et al., 1992; Naudin et al., 1991). Our data support the concept that decreased RYR2 function in cardiac pathology may directly contribute to metabolic inflexibility seen in heart failure. Our data also provide evidence for a role for RYR2 and Ca2+ in regulating lipid utilization and ATP production by transcriptional reprogramming. It is well established that failing hearts switch from metabolizing free fatty acids towards carbohydrate utilization (Stanley, 2005). Knockout and overexpression studies show that changes in transcriptional regulators and enzymes involved in lipid metabolism are sufficient to cause cardiac hypertrophy and steatosis (Haemmerle et al., 2006; Son et al., 2010; Ueno et al., 2008). For example, our data suggest a role of the cytoprotective and metabolism-controlling Sirt1/Foxo1/Pgc1α axis in hearts lacking Ryr2. RYR2 deficiency is sufficient to flip a transcriptional metabolic ‘master switch’, which down-regulates lipid utilization pathways and effectors. We also found reduced insulin signaling and dramatic loss of Glut4, which might be expected to have deleterious effects given the cardioprotective effect of glucose uptake (Liao et al., 2002). Therefore, our study suggests that acute Ryr2 deletion disrupts both lipid and carbohydrate metabolism indirectly via transcriptional programming, as well as directly at the mitochondria. In our previous studies of other cell types, loss of metabolism-pacing Ca2+ communication between RyR channels and mitochondria caused cellular ATP levels to decline and induced the presenilin-dependent expression of key hypoxia response genes (Dror et al., 2008; Liu and O'Rourke, 2008). Similarly, cRyr2KO mice had significantly increased proteins levels of presenilin-1, presenilin-2, HIF1α and HIF1β, and they displayed numerous signs of energy   101 starvation with an associated increase in compensatory mechanisms for energy conservation and optimization. This included decreased expression of uncoupling proteins, downregulated carbohydrate and lipid storage pathways. Thus, cRyr2KO hearts exist in a pseudo-hypoxic stress state. Hypoxia-inducible factors are known to increase Atf3 and decrease Klf15 expression, a combination of events that is sufficient to drive cardiomyopathy, hypertrophy, fibrosis and cell death (Cullingford et al., 2008; Fisch et al., 2007; Huang et al., 2008; Okamoto et al., 2001). While we cannot rule out other parallel mechanisms of cardiac dysfunction in other models of heart failure, our evidence does demonstrate that disruption of RYR2-dependent metabolism is sufficient to cause heart failure. Pharmacological studies have shown that Ca2+ released through RYRs or IP3Rs into adjacent mitochondria modulates apoptosis (Pacher et al., 2002; Tsuboi et al., 2003). When in excess, Ca2+ transfer from SR/ER to mitochondria triggers caspase-3-dependent apoptosis (Luciani et al., 2008). We and others have shown that constitutive intracellular Ca2+ signals are required to stimulate cellular respiration (Rutter et al., 1996) and prevent a hypoxia-like state with calpain-dependent, caspase-3-independent cell death (Dror et al., 2008; Johnson, 2004). Interestingly, calpain-10 is localized to mitochondria (Arrington et al., 2006), where it is poised to sense metabolic state. Our data indicate that disruption of RYR2 function, via its effects on mitochondrial calcium signaling and cellular metabolism, is also sufficient to cause cell death in vivo.  Additional work will be required to further elucidate the mechanisms involved in this atypical cell death mode. We found evidence of autophagy in cRyr2KO hearts similar to the data on cells lacking IP3R (Cárdenas et al., 2010), suggesting that the control of metabolism and autophagy by SR/ER-to-mitochondria Ca2+ shuttling, both the barely perceptible Ca2+ micro-  102 signaling from stochastic channel opening as well as the larger global Ca2+ signals found in excitable cells such as cardiomyocytes, may be a ubiquitous metabolic control system. Our inducible knockout approach was designed to mitigate chronic compensatory gene expression induced by life-long genetic alterations. Accordingly, our data were not confounded by compensation from other ER/SR Ca2+ release channels. We speculate that this may be the reason for the discrepancy between our observations of cardiac hypertrophy and the results obtained in mice with chronic Ryr2 haploinsufficiency, which were protected from pressure overload-induced hypertrophy (Zou et al., 2011). Interestingly, increased levels of cell death were reported in Ryr2+/- mice. The majority of in vivo studies of Ryr2 have used point-mutation knock-in mice, typically with gain-of-function mutations with ‘leaky channels’ (George et al., 2007a; Lehnart et al., 2008). These studies have focused on the susceptibility of these mice to tachycardic arrhythmia. Recently, another gain-of-function knock-in model was created (Ryr2ADA/ADA mice) and reported to undergo calcineurin-independent hypertrophy (Yamaguchi et al., 2011). Additional studies are required to determine whether the hypertrophy observed in our cRyr2KO hearts is mediated through a similar mechanism. Given the fact that basal cytosolic Ca2+ appears to be lower in cRyr2KO cardiomyocytes, one might speculate that a calcineurin-independent pathway is involved. While ours is the first in vivo study to target an intracellular Ca2+ channel and examine metabolism, our conclusions are well supported by more reductionist studies on the roles of RYR2 in cardiac metabolism (Cárdenas et al., 2010; Denton and McCormack, 1990; Giacomello et al., 2010; Glancy and Balaban, 2012; Liu and O'Rourke, 2008), cell survival (Dror et al., 2008; Johnson, 2004; Zou et al., 2011), and cardiac dysfunction (Bidasee et al., 2001; Brillantes et al.,   103 1992; Naudin et al., 1991). In our model, deletion of Ryr2 rapidly led to many alterations in cardiac phenotype. Since our only manipulation was Ryr2 deletion, all the observed changes must ultimately be downstream of RYR2 action in cardiomyocytes, either immediately downstream or indirectly downstream. Future studies are required to elucidate the specific mechanisms involved, and the temporal order in which these events occur.  In summary, our results illustrate that Ca2+ flux through the ER/SR RYR2 Ca2+ channels is required to maintain mitochondrial Ca2+, oxidative metabolism, metabolic transcriptional pathways, and cellular survival in the mouse heart in vivo. These observations, which support concepts advanced from in vitro studies, demonstrate a paramount role for RYR2 in cardiomyocyte energetics. This creates a paradigm in which RYR2 Ca2+ fluxes have simultaneous roles in the regulation of heart rate and rhythmicity (Bround et al., 2012), as well as effects on metabolic mitochondrial pacing, and transcriptional programming in cardiomyocytes. This comprehensive model may explain how functional and metabolic demands of cardiomyocytes can be so exquisitely coupled to ensure energy needs are met.           104 Chapter 5:  Cardiac RYR2 Specifically Promotes Cardiac Glucose Oxidation  5.1   Chapter Summary Cardiac ryanodine receptor (RYR2) Ca2+ release channels and cellular metabolism are both disrupted in heart disease. Recently, we demonstrated that total loss of RYR2 leads to cardiomyocyte contractile dysfunction, arrhythmia, and reduced heart rate. Acute total RYR2 ablation also impaired metabolism, but it was not clear whether this was a cause or consequence of heart failure in this model. Previous in vitro studies have suggested that Ca2+ flux into the mitochondrial can help pace oxidative metabolism, but there is limited in vivo evidence supporting this mechanism. Here, we study heart specific, inducible Ryr2 haploinsufficiency (cRyr2Δ50) mice with a stable 50% reduction in RYR2 protein. This manipulation decreased the amplitude and frequency of cytosolic and mitochondrial Ca2+ signals in isolated cardiomyocytes, without changes in cardiomyocyte contraction. Remarkably, in the context of mildly impaired contractile function in perfused hearts, we observed a decrease in glucose oxidation coupled with an increase in the rate of glycolysis but no change in the rate of fat oxidation. We also observed that cRyr2Δ50 hearts displayed pyruvate dehydrogenase hyper-phosphorylation, indicative of inhibition of this Ca2+-sensitive gatekeeper to glucose oxidation. Metabolomic, proteomic, and transcriptomic analyses revealed additional functional networks associated with altered metabolism in this model. These results demonstrate that RYR2 controls mitochondrial Ca2+ dynamics and plays a specific, critical role in promoting glucose oxidation in cardiomyocytes. Our findings indicate partial RYR2 loss is sufficient to cause metabolic abnormalities seen in heart disease.    105 5.2   Introduction The type 2 ryanodine receptor (RYR2) sarcoplasmic reticulum (SR) Ca2+ release channel plays a central role in cardiac excitation contraction coupling (Lanner et al., 2010). Ca2+ signals generated by RYR2 have also been implicated in heart rate (George et al., 2007a; Monfredi et al., 2013; Yang et al., 2002), hypertrophic gene regulation (Wilkins and Molkentin, 2004; Zou et al., 2011), and cardiomyocyte superstructure (Crossman et al., 2011; Scriven et al., 2013). Partially reduced RYR2 expression, channel density, or signaling have been identified in models of aging (Kandilci et al., 2011), and heart disease (Bidasee et al., 2001; Crossman et al., 2011; Matsui et al., 1995; Milnes, 2001), which are conditions associated with metabolic dysfunction and a lack of energy substrate flexibility (Doenst et al., 2013; Stanley, 2005). However, it remains to be determined whether a partial reduction in RYR2 signalling can be a driver of adult cardiac metabolic dysfunction.  SR Ca2+ release channels are known to transmit privileged Ca2+ signals into adjacent mitochondria (Rizzuto and Pozzan, 2006). Based on in vitro studies, it has been proposed that these Ca2+ signals stimulate mitochondrial oxidative energy metabolism via key tricarboxylic acid (TCA) cycle enzymes (e.g. pyruvate dehydrogenase), the electron transport chain, and the mitochondrial ATPase (Denton, 2009; Glancy and Balaban, 2012; Glancy et al., 2013). These concepts have been extended to cardiac biology where studies have shown a link between mitochondrial Ca2+ and energy production in the heart (Bround et al., 2013; Chen et al., 2012; Gong et al., 2014; Liu and O'Rourke, 2008; Pan et al., 2013). However, direct in vivo evidence that Ca2+ flux from ER/SR specifically paces mitochondrial oxidative glucose metabolism in normally functioning cardiomyocytes has not been published. We recently reported that complete Ryr2 gene knockout in adult mouse cardiomyocytes results in broad defects in   106 metabolism (Bround et al., 2013), but our previous model rapidly progressed to heart failure and cardiac death (Bround et al., 2012) making it difficult to know if mitochondrial metabolism is modulated by RYR2 in normally functioning cardiomyocytes. Given this previous work and the association between RYR2 dysfunction (Bidasee et al., 2001; Crossman et al., 2011; Matsui et al., 1995; Milnes, 2001) and oxidative metabolism in heart disease (Doenst et al., 2013; Stanley, 2005), it is of interest to further explore links between RYR2 and energy production using improved models.  To interrogate the potential links between RYR2 and metabolism in the context of relatively normal cardiac function, we devised a stable model where cardiomyocyte-specific, inducible Cre recombinase was used to delete only one Ryr2 allele (cRyr2Δ50 mice). Here, we report that a stable ~50% loss of RYR2 protein in adult cardiomyocytes causes striking changes in mitochondrial Ca2+ cycling and is sufficient to specifically inhibit oxidative glucose metabolism but not fatty acid oxidation, lactate oxidation or glycolysis. These metabolic effects occurred without significantly affecting cellular contraction or in vivo cardiac output, although we did observe a modest decrease in cRyr2Δ50 heart rate. Our results suggest that pathophysiologically relevant loss of RYR2 can account for the metabolic phenotype of failing hearts and that RYR2 plays a critical role in stimulating glucose oxidation in vivo.       107 5.3   Results First we measured the degree of RYR2 ablation in the cRyr2Δ50 model and determined whether there was compensation from any related Ca2+ channels. We confirmed partial ablation of Ryr2 mRNA and protein in the cRyr2Δ50 mice 3 weeks following the induction of haploinsufficiency with sequential tamoxifen injections which resulted in a ~50% decrease in RYR2 protein levels (Fig. 5.1A-C). Importantly, there was no compensation by other sarcoplasmic reticulum (SR)-Ca2+ release channel genes (Fig. 5.1B).  We next characterized cytosolic and mitochondrial Ca2+ signaling in cardiomyocytes from cRyr2Δ50 mice. We simultaneously measured cytosolic and mitochondrial Ca2+ in isolated cells stimulated with either a single pulse of field stimulation or with periods of continuous 0.5Hz or 6Hz pulses (Fig. 5.2). Both cytosolic and mitochondrial Ca2+ dynamics were significantly different between cells from control and cRyr2Δ50 mice (Fig. 5.2A-C). Specifically, cytosolic and mitochondrial Ca2+ transient systolic amplitudes were diminished in cRyr2Δ50 cardiomyocytes across all stimulation frequencies (Fig. 5.2A-B,D,F). We also observed a striking reduction in the frequency of both cytosolic and mitochondrial Ca2+ oscillations when stimulating cardiomyocytes at 6 Hz (Fig. 5.2C,E,G). Whereas control cardiomyocytes showed overlap between their cytosolic and mitochondrial systolic peaks, cRyr2Δ50 cardiomyocytes displayed a ~10 second lag between peak systolic cytosolic Ca2+ and peak mitochondrial Ca2+ at 6 Hz (Fig. 5.2C). These data suggest that a ~50% RYR2 ablation alters cytosolic Ca2+ signaling and disrupts SR-to-mitochondrial Ca2+ communication.      108  Figure 5.1. Generation of an inducible, heart-specific cRyr2Δ50 knockout model with partial RYR2 ablation. (A) Schematic depicting breeding scheme, experimental design, and analysis timeline for cRyr2Δ50 mice. (B) cRyr2Δ50 cardiomyocytes have a specific reduction in Ryr2 mRNA, and no compensation from other known ER/SR Ca2+ channels (n = 4-8, *p ≤ 0.05). Blue bars = Control, Purple bars = cRyr2kΔ50 throughout. (C) Cardiac RYR2 protein levels in cRyr2Δ50 mice 3 weeks after tamoxifen (n = 7, *p ≤ 0.05). All data plotted as mean ± SEM. Control = Ryr2flox/wildtype + tamoxifen; cRyr2Δ50: Ryr2flox/wildtype x Mhy6-MerCreMer+ + tamoxifen.    109                   Figure 5.2. Altered cytosolic and mitochondrial Ca2+ homeostasis in cRyr2Δ50 cardiomyocytes. (A-C) Representative simultaneous Fura-2 and Rhod-2 fluorescence traces from control and cRyr2Δ50 cardiomyocytes. Continuous recordings (A-C) are of a single cell per treatment group. Fura-2 ratio is plotted as a blue line and Rhod-2 intensity is plotted as a red line in units relative to baseline (RU). Scale bar is 0.1 RU of reporter signal in the vertical axis and 20 s in the horizontal axis. Representative Fura-2 and Rhod-2 traces during pulse (A) and 0.5Hz continuous (B) field stimulation (grey triangles). Scale axis begins at 0.9 RU. Inset graphs show average peak systolic Fura-2 ratio and Rhod-2 intensity relative to baseline. (C) Representative Fura-2 and Rhod-2 traces during 6 Hz continuous field stimulation (grey triangle). Scale axis begins at 8.5 RU. (D) Average Fura-2 ratio in control and cRyr2Δ50 cardiomyocytes during 6 Hz stimulation. White bars = control, solid color bars = cRyr2Δ50 throughout (mean ± SEM). Sys-Base denotes the difference between average peak systolic and baseline measurements, Dias-Base denotes the difference between average minimal diastolic and average baseline measurements, and Sys-Dias   110 denotes the difference between peak systolic and minimum diastolic measurements. (E) Average frequency of cytosolic Ca2+ transients elicited during 6 Hz stimulation. (F) Average Rhod-2 intensity in control and cRyr2Δ50 cardiomyocytes during 6 Hz stimulation. (G) Average frequency of elicited mitochondrial Ca2+ transients observed during 6 Hz stimulation. Control n = 63 cells, cRyr2Δ50 n = 141 cells, 3 independent cell preparations form 3 mice per treatment;  *p ≤ 0.05. All data plotted as mean ± SEM. Control = Ryr2flox/wildtype + tamoxifen; cRyr2Δ50: Ryr2flox/wildtype x Mhy6-MerCreMer+ + tamoxifen.  We next examined the function of cRyr2Δ50 cardiomyocytes, in vitro and in vivo. We did not observe significant changes in diastolic length, fractional shortening, the time to 50% peak contraction, or the time to 50% relaxation (Fig. 5.3A-D), indicating that contractile function in individual cells is normal. Cardiac function in isolated perfused working hearts revealed a decrease in cardiac output, rate pressure product, and cardiac work in cRyr2Δ50 hearts (Fig. 5.3E-G). However, these changes were modest compared to the dramatic mechanical dysfunction observed in total Ryr2 knockout hearts (Bround et al., 2013). Echocardiography also failed to show any significant differences in heart function in cRyr2Δ50 mice 3 or 20 weeks following tamoxifen injection (Fig. 5.3H-I). We have previously demonstrated that complete Ryr2 gene knockout results in reduced heart rate and fatal arrhythmia (Bround et al., 2013). In the present study, we found an intermediate level of bradycardia in cRyr2Δ50 mice using subcutaneously implanted ECG telemetry in freely moving, un-anesthetized mice (Fig. 5.3J-K). This suggests that ~50% RYR2 ablation is sufficient to depress heart rate in the absence of cardiac mechanical dysfunction. Collectively, these experiments suggest that a stable, ~50% reduction in RYR2 is compatible with well-preserved function in vivo and in vitro, although a mild impairment can be identified in the ex vivo working heart system.      111   Figure 5.3. Cardiac function and heart rate in cRyr2Δ50 mice. Isolated cardiomyocytes were assessed for (A) average diastolic length and (B) fractional shortening upon stimulation (n = 3;  *p ≤ 0.05.). Isolated cardiomyocyte contraction rate reported as (C) time to 50% peak contraction and (D) time to 50% peak relaxation. White circles = control, Black circles = cRyr2Δ50. (E) Cardiac output, (F) rate pressure product, and (G) cardiac work measured during working heart perfusions (Control n = 10, cRyr2Δ50 n=13; *p ≤ 0.05.). White bars = control, Black bars = cRyr2Δ50; throughout. Echocardiograms of control and cRyr2Δ50 mice 3 weeks following tamoxifen treatment. (H) Average cardiac output and (I) fractional shortening of control and cRyr2Δ50 mice 3 and 20 weeks following tamoxifen treatment (n =5;  *p ≤ 0.05.). (J) Average heart rate from implantable ECG radio telemetry Blue points denote days prior to tamoxifen injections (red points), but following surgical recovery and the removal of analgesics). Heart rate is normalized to the average heart rate measured during baseline (dashed line). Red dashed line denotes average heart rate of the total cRyr2KO mice reported in our previous publication (Bround et al. 2012, Cardiovascular Research) (K) Average heart rate before and after tamoxifen treatment (mean ± SEM; n = 5;  *p ≤ 0.05). All data plotted as mean ± SEM. Control = Ryr2flox/wildtype + tamoxifen; cRyr2Δ50: Ryr2flox/wildtype x Mhy6-MerCreMer+ + tamoxifen.   112  We next assessed cardiac energy metabolism in functional cRyr2Δ50 cardiomyocytes. Unlike the total RyR2KO hearts (Bround et al., 2013), there was no change in total ATP levels in cRyr2Δ50 hearts (Fig. 5.4F) suggesting that they are not significantly energy deprived but rather that energy metabolism is sufficient to sustain adenine nucleotide energy charge. Dynamic metabolic measurements of perfused working cRyr2Δ50 hearts were employed to assess utilization and oxidation of multiple energy substrates. Remarkably, during simultaneous measurements in the same hearts, we observed a significant decrease in glucose oxidation and an increase in glycolysis (Fig. 5.4A,B). Fatty acid oxidation and lactate oxidation were not altered in these cRyr2Δ50 hearts (Fig. 5.4C,D). A metabolomic survey of tricarboxylic acid cycle intermediates in cRyr2Δ50 heart tissue displayed significant decreases in fumarate and α-ketoglutarate, as well as a general trend towards decreased levels of other tricarboxylic acid cycle intermediates (Fig. 5.4E). Glycolytic metabolites upstream of entry into the tricarboxylic acid cycle, including pyruvate, remained unchanged in cRyr2Δ50 hearts (Fig. 5.4E). These data demonstrate that glucose utilization is altered in cRyr2Δ50 hearts in the absence of major cardiac dysfunction and suggest that RYR2 has a direct and specific role in promoting the full oxidation of glucose in cardiac tissues in vivo. To examine the molecular mechanism involved in this metabolic shift, we first focused on PDH, the enzyme that catalyses the conversion of pyruvate to acetyl-CoA. PDH is known to be a critical gatekeeper between glycolysis and complete glucose oxidation (Patel et al., 2014), and its activity is regulated by complex mechanisms including inhibitory phosphorylation at three sites, including serine-293. PDH phosphorylation is mediated by a family of PDH kinases (PDK 1-4) while its dephosphorylation is catalyzed by a Ca2+ sensitive phosphatase (Denton, 2009).   113 PDH phosphorylation was significantly increased in the hearts of cRyr2Δ50 mice fasted over-night (Fig. 4G), but not in un-fasted hearts (data not shown). This is reminiscent of the conditions required to uncover changes in PDH regulation in mice lacking the mitochondrial Ca2+ uniporter (Pan et al., 2013). The increased PDH phosphorylation is consistent with our observed specific inhibition of glucose oxidation in cRyr2Δ50 hearts (Fig. 5.4B). Transcript levels of Pdk1, Pdk2, Pdk3, and Pdk4 were not significantly changed in cRyr2Δ50 hearts (Fig. 5.4H). However, we observed relative decreases in PDK1 and PDK4 in the proteome of mouse hearts collected during basal conditions (Fig. 5.6B). These observations suggest that increased PDH phosphorylation is most likely explained by decreased activity of PDP phosphatase, despite a compensatory reduction in expression of PDK1 and PDK4 activity in cRyr2Δ50 hearts (see below). While these results suggest that PDH regulation is highly reliant on metabolic context, our findings indicate RYR2 plays a key role in PDH activation.               114  Figure 5.4. Metabolomic and working heart analyses of cRyr2Δ50 mice reveal a specific defect in glucose oxidation, associated with hyper-phosphorylation of PDH. (A) Non-oxidative glycolysis and (B) complete glucose oxidation was simultaneously measured in working hearts (control n = 5, cRyr2Δ50 n = 6;  *p ≤ 0.05). Light blue bars = control, purple bars = cRyr2Δ50 throughout. At the same time, (C) palmitate oxidation and (D) lactate oxidation were measured (control n = 5, cRyr2Δ50 n = 7; *p ≤ 0.05). (E) Diagram showing the results of targeted mass spectroscopy-based metabolomic analysis of TCA Cycle acid intermediaries and glycolysis pathway sugar intermediates (n = 8;  *p ≤ 0.05; grey = unmeasured). (F) Average heart ATP, ADP, and AMP as measured by HPLC-chromatography on whole homogenized hearts (n = 3;  *p ≤ 0.05). (G) Pyruvate Dehydrogenase Kinase E1-alpha subunit serine 293 phosphorylation levels in mice fasted overnight (Ctrl n=5, cRyr2Δ50 =6; *p ≤ 0.05). (H) Pyruvate dehydrogenase kinase isoform mRNA levels of in cRyr2Δ50 hearts (control n = 5, cRyr2Δ50 n = 4, *p ≤ 0.05). All data plotted as mean ± SEM. All data plotted as mean ± SEM. Control = Ryr2flox/wildtype + tamoxifen; cRyr2Δ50: Ryr2flox/wildtype x Mhy6-MerCreMer+ + tamoxifen.    115 In an effort to provide unbiased insight into the functional and metabolic effects of partial RYR2 reduction, we performed proteomic analysis to compare extracts from control hearts and cRyr2Δ50 hearts, spiked with known amounts of SILAM wildtype heart tissue to allow quantitative comparisons (Fig. 5.5A) (Zanivan et al., 2012). With this approach, we identified 1755 unique proteins with high-confidence (Fig. 5.5B), and analyzed 1702 proteins after manually curating-out blood-borne protein contaminants. The arbitrarily chosen 35 most highly increased proteins, based on fold change, included: Sacm1l, a regulator of IP3 signaling; Bmp10, a key regulator of heart development; atrial natriuretic factor, a cardio-protective hormone and biomarker of cardiac dysfunction (Song et al., 2015); and Ahnak2, a large t-tubule protein which interacts with RYR2 (Komuro et al., 2004) (Fig. 5.5B). On the other hand, the 35 most highly decreased, based on fold change, proteins included: Homer, which interacts with RYR2 to decrease its open probability (Pouliquin et al., 2009), Akt, a central protein kinase in insulin signaling; and RYR2. Network analysis identified clusters of interacting proteins involved in heart muscle contraction, actin binding/motility, and muscle cell differentiation that were increased (Fig. 5.5C), as well as clusters of proteins involved in fatty acid oxidation/metabolism, mitochondrial inner membranes/matrix, contractile fibers and TCA cycle that were decreased in the cRyr2Δ50 heart proteome (Fig. 5.5D). This unbiased survey illustrates that hearts undergo a substantial degree of metabolic and contractile remodeling as a result of partial Ryr2 deletion.      116                     Figure 5.5: Proteomic analysis of cRyr2Δ50 hearts. (A) Schematic depicting the experimental design of the proteomics experiment, including spike-in with stable isotope-labeled heart tissue (SILAM hearts) to enable quantitative normalization. Four mice per treatment group were pooled and analyzed simultaneously by mass spectroscopy. (B) Distribution relative protein abundance measurements, with the 25 most enriched proteins (green) and 25 most depleted proteins (red) between treatment groups listed. Analysis of up-regulated (C) and down-regulated (D) proteins identified significantly enriched networks (FDR < 0.05)    117                     Figure 5.6: Parallel analysis of functional protein categories in cRyr2Δ50 hearts. Visualization of the changes of key cardiac proteins arranged into functional clusters of (A) excitation-contraction coupling proteins, (B) glycolysis and glucose oxidation proteins, or (C) fat oxidation and other key metabolic proteins. Data is plotted as the fold difference between cRyr2Δ50 and control hearts (one sample per group made with four pooled hearts per sample to reduce variation). Red ≤ 0.75 fold change, orange ≤ 0.85 change, pale green ≥ 1.15 enrichment, and dark green ≥ 1.25 enrichment. Grey schematics show location of presented glucose and fat oxidation proteins in their respective pathways.    118                   Figure 5.7: The Proteome of the electron transport chain, ATP synthase complex, and other metabolic effectors in cRyr2Δ50 hearts. Visualization of the changes of additional cardiac proteins arranged into functional clusters of electron transport chain, metabolic, calcium sensitive, and cardiac pathology associated proteins. Data is plotted as the fold difference between cRyr2Δ50 and control hearts (one sample per group made with four pooled hearts per sample to reduce variation). Red represents a ≤ 0.75 fold change, orange a ≤0.85 change, pale green ≥ 1.15 fold enrichment, and dark green ≥ 1.25 fold enrichment.      119 We used the proteomic data to assess the status of cRyr2Δ50 cardiomyocyte sarcomeres and mitochondria (Fig. 5.6, 5.7). This targeted survey of core excitation-contraction proteins identified a modest decrease in the Na/Ca2+ exchanger NCX1 (Fig. 5.6A). We also observed modest decreases in the troponin/tropomyosin complex and a large increase in atrial isoforms of regulatory and essential myosin light chains (MLCs) which have been associated with heart disease (Hernandez et al., 2007), as well as increased levels of calponins, proteins that interact with actin and tropomyosin which, when overexpressed, can partially rescue hearts from dilated cardiomyopathy (Lu et al., 2014). A survey of the glycolytic and glucose oxidation pathways showed no changes in core pathway enzymes (Fig. 5.6B). Instead, we observed reductions only in FBP2, an anabolic protein that counteracts the activity of PFK1, the key gatekeeper of glycolysis (Mor et al., 2011), as well as PDK1 and PDK4, inhibitory kinases of the PDH glucose oxidation gatekeeper (Fig. 5.6B) (Patel et al., 2014). These data suggest a reduction of inhibitory signals to both glycolysis and glucose oxidation in cRyr2Δ50 hearts despite the reduced glucose oxidation and context dependent PDH hyper-phosphorylation. A similar survey of the fat oxidation pathway shows modest decreases in the carnitine acyl-carnitine transporter and some β-oxidation proteins (Fig. 5.6C). However, since these changes failed to result in reduced palmitate oxidation rates, these changes must be within the range of dynamic regulation in this model. We also observed a large decrease in NAD kinase and a large increase in nicotinamide N-methyltransferase (NNT), changes that are consistent with the cell trying to preserve reducing equivalents for the TCA cycle but which may also compromise the antioxidant capacity of the cell (Sheeran et al., 2010) (Fig. 5.6C). We did not observe robust changes in the ETC or ATP synthase machinery sub-proteomes (Fig. 5.7). Collectively, this proteomic analysis   120 revealed changes initiated by the stable 50% reduction in RYR2 that are likely to underlie effects on metabolism. Our mass spectrometry-based proteomic analysis was not able to assess every gene product, despite the ability to simultaneously examine 1702 heart proteins in a highly quantitative manner. Thus, RNA sequencing was also employed to provide genome-wide analysis of changes in cRyr2Δ50 hearts. There were relatively few changes in mRNA levels in the transcriptome of cRyr2Δ50 hearts that reached genome-wide significance (Fig. 5.8A). However, notable differences included decreased expression of Ryr2 (Fig. 5.8B), voltage gated Na+ channel Scn4b (Fig. 5.8C), and mylk4 (Fig. 5.8D), a novel isoform of myosin light chain kinase that is down regulated in heart failure (Herrer et al., 2014). We also found decreased expression of Ucp1 and Cbr2 genes that utilize TCA reducing equivalents for non-energy producing processes but potentially at the expense of protection from reactive oxygen species (Hoerter et al., 2004) (Fig. 5.8E). Biosynthetic genes were also significantly decreased in cRyr2Δ50 hearts, including key gluconeogenesis gene Pck1 and Gpd1, a protein that scavenges glycolysis products for sugar biosynthesis (Fig. 5.8F). Cbr2 and Gpd1 were also observed to be decreased in the heart proteome (Fig. 5.6C, 5.7). We also observed a general up-regulation of extracellular structural and remodeling genes consistent with cardiac pathology (de Haas et al., 2014) (Fig. 5.8G). In summary, the cRyr2Δ50 heart transcriptome also revealed some degree of contractile and metabolic modulation.       121  Figure 5.8: Transcriptomics analysis of cRyr2Δ50 hearts. (A) Biological Gene Ontology (PANTHER) groups of significantly changed genes in the cRyr2Δ50 heart transcriptome. Changes in the mRNA levels of (B) Ryr2, (C) excitation contraction, (D) core contractile, (E) core energetic, (F) biosynthetic, and (G) extracellular matrix genes in cRyr2Δ50 hearts (n=4, *p ≤ 0.05). All data plotted as mean ± SEM. All data plotted as mean ± SEM. Control = Ryr2flox/wildtype + tamoxifen; cRyr2Δ50: Ryr2flox/wildtype x Mhy6-MerCreMer+ + tamoxifen.          122 5.4   Discussion The goal of the present study was to examine the effects of ~50% reduction in RYR2 on heart metabolism and function. We found that a stable 50% loss of RYR2 protein impaired mitochondrial Ca2+ signaling, reduced Ca2+-dependent PDH activation, and specifically reduced oxidative glucose metabolism. Importantly, these changes occurred in the absence of robust mechanical dysfunction, meaning that they are primary defects downstream of RYR2. Unbiased, proteomic and transcriptomic surveys of cRyr2Δ50 hearts revealed alterations in contractile and metabolic gene networks predicted to increase glucose oxidation and RYR2 Ca2+ release, suggesting mechanisms for compensation in this model.  Our principle finding is that RYR2 specifically promotes glucose oxidation in cardiomyocytes. To the best of our knowledge, this is the first in vivo demonstration of an ionic mechanism that specifically controls glucose oxidation, but not fat oxidation. A wealth of in vitro research has shown that mitochondrial Ca2+ signaling is important for the oxidation of energy fuel substrates by stimulating TCA cycle enzymes, the ETC, and ATP synthase (Denton, 2009; Glancy and Balaban, 2012; Glancy et al., 2013). Specifically, it has been previously proposed that glucose oxidation is regulated by Ca2+ signaling through the key gatekeeper for pyruvate entry into the TCA cycle, PDH, which is activated by a Ca2+ sensitive phosphatase (PDP) (Denton, 2009; Patel et al., 2014). Our findings demonstrate that RYR2 is upstream of mitochondrial Ca2+ signals, affects PDH phosphorylation levels, and has a specific role in promoting glucose oxidation. The finding that RYR2 plays a specific and sensitive role in promoting glucose oxidation may represent a key mechanism by which the metabolic demands of excitation-contraction are directly coupled to the rate of ATP production in cardiomyocytes.   123 Glucose oxidation contributes to cardiomyocyte energy metabolism and is preferentially up-regulated in periods of oxidative stress or increased metabolic demand (Stanley, 2005). It is notable that we only observed effects of RYR2 haploinsufficiency on PDH phosphorylation in fasted cRyr2Δ50 mice. Fasting is a situation that is predicted to increase the expression of PDK proteins, which would increase the inhibitory phosphorylation tone on the PDH system (Jeong et al., 2012). In this situation, the importance of the phosphatase activity of PDP may become critical in reactivating PDH to allow for effective glucose oxidation during metabolic stress (Sun et al., 2015). Our findings suggest that, in this context, RYR2 becomes necessary to fully drive PDP activity and prevent hyper-phosphorylation of PDH. The effects of RYR2 on PDH activation may have been partially obscured by basal reductions in PDK1 and PDK4 in the cRyr2Δ50 heart proteome, suggesting the possibility that PDH activation may normally be even more sensitive to RYR2 than our experiments revealed. Notwithstanding, our data clearly demonstrate, for the first time, that RYR2 is critically involved in preferentially increasing glucose oxidation during metabolic stress. Our previous research measured the effects of total RYR2 deletion on oxidative metabolism and reported reduced heart ATP levels and a general reduction in total oxidative energy metabolism (Bround et al., 2013). The present study demonstrates that a 50% RYR2 reduction specifically disrupts glucose oxidation, which suggests there may be a hierarchy where oxidative metabolism of multiple substrates is generally Ca2+ sensitive, but glucose oxidation is more attuned to changes in Ca2+ homeostasis. Since glucose oxidation and glycolysis are frequently uncoupled in models of heart disease before the general impairment of oxidative metabolism seen in heart failure (Doenst et al., 2013; Stanley, 2005), these data also suggests that progressive   124 RYR2 dysfunction in disease may contribute to mounting metabolic dysfunction in a ‘dose-dependent’ manner. An interesting finding of our metabolic studies is that glucose oxidation was decreased while lactate oxidation levels remain unchanged in our cRyr2Δ50 hearts. This is somewhat unexpected because both lactate oxidation and glycolysis generate pyruvate as a step towards full oxidation. As such, both lactate and glucose oxidation rates are dependent on PDH activity, and in theory, should be affected similarly if RYR2 is required for PDH complex activation. This creates the possibility that reductions in glucose oxidation rates are not due to decreased pyruvate entry into the TCA cycle, but rather that glycolytically-derived pyruvate is being preferentially shunted towards other biochemical fates. We believe that pyruvate carboxylation for anaplerosis or biosynthesis is unlikely since we observe a general depletion of TCA intermediaries as well as a down regulation of key biosynthetic genes. We think it is more likely that glycolytically-derived pyruvate is being converted to lactate to resupply the cytosolic NAD+ supply to support increased glycolysis rates. The apparent disconnect between lactate and glucose oxidation rates likely reflects the metabolic compartmentalization of glycolytically-derived lactate and exogenous lactate (Chatham et al., 2001), as well as the observation that lactate has a 10-40% higher efficiency as an oxidative substrate than exogenous pyruvate (Brooks et al., 1999; Molé et al., 1978). We would posit that in the cRyr2Δ50 model, there is decreased PDH activity in some contexts and that this manifests itself to a greater extent in glucose oxidation rates due to the substrate efficiency differences between lactate and glycolytically-derived pyruvate. However, we cannot currently discount that a portion of the glucose oxidation decrease may be due to upregulated glycolysis rates.    125 Our results indicate that a 50% reduction in RYR2 changes the frequency and amplitude of cytosolic and mitochondrial Ca2+ signals. Since we did not observe major alterations in the levels of other Ca2+ cycling proteins, our data suggest that altered Ca2+ signalling is due to changes in RYR2 expression. Since groups of RYR2 normally function in concert, due to proximity and physical interaction (Scriven et al., 2013), it is possible that decreasing RYR2 abundance reduced the functional coupling within the RYR2 signalling apparatus leading to more gradual activation and deactivation of Ca2+ transients. Our data clearly illustrate the disrupted privileged communication between the SR and mitochondria. Mitochondria are highly impermeable to Ca2+ and rely on microdomain signaling where the mitochondrial Ca2+ uniporter is paired with Ca2+ release channels to drive mitochondrial Ca2+ uptake (Rizzuto and Pozzan, 2006). Our data suggest that when RYR2 is reduced, Ca2+ cannot flow as quickly between these two organelles via microdomain signaling. It is likely that deleting 50% of RYR2 proteins disrupts SR-mitochondria signaling structures, since RYR2 is the primary Ca2+ channel of these signaling complexes and because RYR2 may also play a structural role in mitochondrial tethering (García-Pérez et al., 2011; Min et al., 2012). Analysis of the role of RYR2 in cardiomyocyte ultra-structure using this mouse model could form the basis of an interesting future study. Another important finding of this study is that a 50% decrease in RYR2 levels is sufficient to significantly reduce heart rate in cRyr2Δ50 mice. In our previous work on total Ryr2 knockout mice (cRyr2KO), we noted a substantial reduction in heart rate, as well as tachycardic arrhythmias (Bround et al., 2012). Our results here show that even in the absence of acute mechanical dysfunction, stable Ryr2 haploinsufficiency is sufficient to reduce heart rate which provides evidence that this is a bona fide effect of RYR2 signalling and not a consequence of heart dysfunction observed in the total cRyr2KO model. The decrease in heart rate we observed   126 in the cRyr2Δ50 mice was intermediate to the reduction we observed in the cRyr2KO model, suggesting that the stimulatory effect of RYR2 on heart rate is dose dependent (Bround et al., 2012). Together, these findings provide strong evidence that RYR2 plays a critical role in cardiac pace-making and supports a model where heart rate is regulated by an ensemble of SR and plasma membrane ionic channels (Monfredi et al., 2013). The cRyr2Δ50 model is a significant improvement on previous loss-of-function RYR2 models as it is not lethal and does not show the dramatic changes in cardiac mechanical function found in total cRyr2KO mice. This provides a better model with which to address phenomena that are directly downstream of RYR2. The cRyr2Δ50 model is also valuable because it greatly reduces the potential for confounding long-term compensatory effects of global, life-long mutations (Zou et al., 2011), as well as the putative period of tamoxifen drug-effects observed in some studies (Koitabashi et al., 2009). Additionally, induced, haploinsufficiency models a situation analogous to disease models which observe either a decrease in RYR2 expression or function similar to the levels in our model (Bidasee et al., 2001; Crossman et al., 2011; Kandilci et al., 2011; Matsui et al., 1995; Milnes, 2001). Thus, the cRyr2Δ50 model is a valuable tool to study how RYR2 dysfunction contributes to human health and disease. Numerous studies have reported reduced RYR2 levels or function in diabetic cardiomyopathy (Bidasee et al., 2001), heart failure (Matsui et al., 1995; Milnes, 2001), and aging (Kandilci et al., 2011) implicating RYR2 dysfunction as an element of heart disease and its pre-disposing conditions. Heart disease is also associated with reductions in the oxidative capacity of the heart and reductions in the metabolic flexibility including reduced glucose oxidation and increased glycolysis (Stanley, 2005). This study shows that a chronic, 50% reduction of RYR2 in adult   127 animals, similar to what is seen in these disease models, is sufficient to disrupt normal glucose oxidation in cardiomyocytes. This suggests that RYR2 dysfunction may contribute to the impaired metabolic flexibility seen in heart disease.     128 Chapter 6:  Conclusion 6.1   Summary The research presented in this thesis seeks to determine which cardiomyocyte cellular processes are downstream of RYR2 channel signaling. Using a tissue specific, inducible gene knockout mouse model, we determined that Ryr2 deletion leads to substantially reduced cardiac contraction, as well as cardiac hypertrophy, cardiomyopathy, and death (Bround et al., 2012). This provides compelling confirmatory evidence that RYR2 is needed for excitation contraction coupling (ECC) in adult animals and suggests that loss of Ryr2 can be a driver of cardiac pathology. We also observed that cRyr2KO animals displayed a significant reduction in heart rate and incidences of cardiac tachycardic arrhythmia (Bround et al., 2012),  providing evidence that RYR2 plays a non-redundant role in cardiac pacemaking and rhythm. Further studies using the cRyr2KO model revealed that Ryr2 knockout lead to a general reduction in mitochondrial ATP production, and that this was associated with the induction of cell death associated with calpain 10 activation (Bround et al., 2013). This provides evidence that RYR2 is a significant driver of cardiac oxidative metabolism in vivo and that the Ca2+ channel plays a role in inhibiting mitochondrial activated cell death. However, because we observed such profound cardiac dysfunction in this model, it remains possible that observed reductions in heart rate and metabolism were consequences of heart failure instead of reduced RYR2 signaling. To determine whether reducing RYR2 expression changes heart rate and metabolism in the absence of heart failure, we also studied a tissue specific, inducible 50% knockdown model where we deleted only one Ryr2 allele. With these cRyr2Δ50 mice we were able to induce a stable, 50% reduction in RYR2 protein in cardiomyocytes. In this context we did not observe severe defects in heart function or contractility in individual cardiomyocytes, nor did we see   129 progression to heart failure and mortality in experimental animals. We also did not observe gross decreases in cellular ATP levels. We did however note that cRyr2Δ50 hearts displayed a moderate reduction in heart rate, intermediate to what we observed in cRyr2KO animals suggesting that the effects of reducing RYR2 on heart rate persist in the absence of gross cardiac dysfunction. This provides compelling evidence that RYR2 does play a non-redundant role in cardiac pacemaking. In addition we observed that induced Ryr2 haploinsufficiency did not result in a general decrease in oxidative metabolism, but instead displayed a specific defect in glucose oxidation. This decrease was associated with altered mitochondrial Ca2+ uptake and a context dependent hyperphosphorylation of the pyruvate dehydrogenase complex (PDH), suggesting that RYR2 plays a role in the activation of this important metabolic enzyme. Collectively, this work shows that RYR2 has a specific role in promoting glucose oxidation even in the context of relatively normal heart function.  6.2   General Caveats Before treating the individual areas of this thesis, there are some general considerations, caveats, and limitations to this research that require discussion. A major confounding factor in this research is the cardiac pathology that accompanies Ryr2 gene knockout. I would argue that ultimately, all of the changes we observe in cRyr2KO cardiac biology are downstream of Ryr2 deletion, since that was our sole experimental manipulation in this model. However, it is impossible to conclude which aspects of the cRyr2KO model phenotype result from a direct effect of RYR2 ablation, and which might be an indirect consequence of reduced cardiac function. This means that the cRyr2KO studies suggest mechanistic links between RYR2 and the effects observed, rather than proving direct signaling   130 interaction. To determine which aspects of heart biology are directly linked to RYR2 signaling we utilized the cRyr2Δ50 model, where we were able to study the effects of reduced RYR2 without the confounding effects of heart failure. This allowed us to conclusively show the involvement of RYR2 in promoting glucose oxidation and maintaining normal heart rate. There, however, remain aspects of the cRyr2KO studies which we have not yet observed or recapitulated in the cRyr2Δ50 model, including cardiac hypertrophy, arrhythmia, and cell death. We did not perform experiments to specifically search for these effects, choosing instead to focus on the role of RYR2 in cardiac metabolism. While future studies may uncover these phenomena in the cleaner cRyr2Δ50 model, at this point I cannot conclusively say whether they are a direct result of RYR2 ablation or a downstream consequence of heart failure. Therefore, the role of RYR2 in hypertrophy and cell death remain probable targets of RYR2 signaling, while the metabolic and heart rate effects of RYR2 are conclusively affected by the Ca2+ channel. Another major consideration is the use of tamoxifen as a driver of inducible cre/lox gene deletion technology.  While we controlled for any potential cardiac effects of tamoxifen in our studies by using tamoxifen injected controls throughout, it has been reported that in some cardiac models the specific association of tamoxifen with inducible cre is somehow cardiotoxic, possibly due to disruptive effects caused by DNA recombination events themselves (Koitabashi et al., 2009). Since this is an unavoidable aspect of our approach, and not something that can be trivially controlled for, it may have manifested in our experiments. It has been suggested that either using a lower dose tamoxifen drug regimen over a series of weeks or waiting 21 days following an intraperitoneal tamoxifen drug regimen are good strategies to avoid cardiotoxicity (Koitabashi et al., 2009). However, given that Ryr2 gene deletion is embryonic lethal (Takeshima et al., 1998), we were forced to utilize an inducible gene deletion approach.   131 Moreover, since induced Ryr2 deletion in adult animals resulted in a rapid onset heart failure and death in most experimental animals within 10 days, both the ‘low and slow’ and ‘wait it out’ approaches for tamoxifen deletion were incompatible with our cRyr2KO studies. As such, it is possible that some tamoxifen/cre-lox effects persist in the cRyr2KO model studies. We did account for any putative tamoxifen driven recombination toxicity in our cRyr2Δ50 model by waiting one month following tamoxifen injections to avoid any potential drug effects. This means that the cRyr2Δ50 experiments benefit from removing both the confounding effects of hear failure and the potential effects of our tamoxifen driven gene deletion. A general feature of this thesis is the difference in phenotype between the cRyr2KO and cRyr2Δ50 models. On the surface, this would seem intuitive: the knockout model has a complete loss of the Ryr2 gene, while the haploinsufficiency model retains one functional allele. However, at the time points we considered, both the cRyr2KO and cRyr2Δ50 models display ~50% reduction in RYR2 protein, yet had remarkably different phenotypes. Therefore, a question is why does a 50% reduction of protein in a gene deletion model lead to heart failure, while a 50% reduction in protein in what is effectively a knockdown model lead to a much milder phenotype. I think this has to do with the fundamental differences between inducible knockout and knockdown studies. It has long been noted that lifelong, constitutive gene knockouts are often extensively compensated for and that inducible gene changes often uncover much more robust phenotypes (De Souza et al., 2006; Rossi et al., 2015). Indeed, constitutive Ryr2 haploinsufficiency bears remarkably little phenotype in both mice and rabbits (Alvarado et al., 2014; Zou et al., 2011), suggesting considerable compensation in these models and showing the value of my inducible approach. What is less known, is the difference between inducible gene knockouts and knockdowns; mechanistically what differs between a stable reduction in protein   132 levels and the gradual degradation and complete loss of that same protein. In this case I can only speculate, but I imagine the difference may stem from the capacity of cRyr2Δ50 mice to produce new RYR2 channels and to turnover damaged or dysfunctional RYR2. In this way the cRyr2Δ50 model may posses a fully functional, perhaps even optimized, complement of 50% RYR2 expression which is apparently enough to maintain normal heart function. The cRyr2KO model, conversely, cannot adequately replace RYR2 molecules, and as a result may represent an uncompensated for reduction and/or the accumulation of damaged RYR2. It is an interesting problem. I do not think this is a critical flaw with these studies, but it persists as an interesting question that this data does not fully explain.  6.3   RYR2 and Excitation Contraction Coupling The current model of excitation-contraction coupling predicts a non-redundant role for RYR2 in activating the contractile machinery in cardiomyocytes. Our results strongly support this model:  we saw that Ryr2 knockout rapidly and significantly diminished the heart’s ability to contract and generate cardiac output. Given the strength and depth of the literature supporting the role of RYR2 in this process, this was not a surprising finding (Bers, 2002; Lanner et al., 2010). Although, it should be noted that targeted deletion of NCX in adult cardiomyocytes showed no deleterious effects on cardiac contraction (Henderson et al., 2004; Pott et al., 2005; 2007), an unexpected finding that showcases the value of  re-testing core theories using new approaches and targeted gene-deletion studies. Furthermore, since SR Ca2+ release is diminished in heart failure while plasma membrane Ca2+ cycling via upregulation of the L-type channel and NCX is significantly increased (Balke and Shorofsky, 1998; Bers, 2006; Gomez, 1997), it was worth testing whether RYR2 deletion could be tolerated by extensive and heretofore unprecedented   133 compensation. However, my results clearly show that RYR2 is critical for ECC and required for heart function and survival as the current paradigm predicts.  What may be an interesting finding from this area of the thesis is that we did not see any obvious reductions in cardiac function and cardiomyocyte sarcomere shortening in the cRyr2Δ50 model. This would seem to indicate that in a stable, chronic situation a 50% reduction in RYR2 is well tolerated by the heart.  This suggests that only 1/2 of the RYR2 channels are required to maintain normal cardiac function and that the heart expresses significantly more RYR2 than it needs. This idea, that the heart has far more RYR2 than it theoretically needs, has long been suggested by theoretical models (Bers, 2001), but these results provide in vivo evidence that this is indeed the case. A caveat of our ECC findings is that there are some key mechanistic differences in excitation-contraction coupling between organisms. Most critical for this thesis are the differences between humans and rodents (Bers, 2002). In humans, it is believed that ~70% of the total Ca2+ flux in ECC coupling is provided by SR Ca2+ stores and RYR2, with the remaining ~30% originating from plasma membrane Ca2+ signaling (Bers, 2002). Rodents, meanwhile, rely on SR Ca2+ signaling for >90% of their ECC needs (Bers, 2002). This means that rodent studies may slightly bias towards the importance of SR Ca2+ and the role of RYR2 in ECC and other cardiac processes. This creates the possibility that in human biology reductions in RYR2 may be better tolerated or perhaps compensated for in ways we were unable to detect using our mouse models. This is an unavoidable caveat of using rodent models in cardiac research.      134 6.4   RYR2 and Heart Rate The data presented in this thesis provides compelling evidence that RYR2 is a critical component of cardiac pacemaking. We found that complete deletion of Ryr2 in the heart decreased heart rate by ~100 BPM (~20%), which persisted from the time of gene deletion to animal death. Additionally we found that induced Ryr2 haploinsufficiency caused heart rate to decrease by 50 BPM (~10%), demonstrating that even in the context of well perserved cardiac contraction, decreases in RYR2 levels result in significant decreases in basal heart rate. Interestingly, the amount heart rate decreased in the cRyr2Δ50 model was intermediate to the drop in heart rate observed in the cRyr2KO model, suggesting that RYR2 has a dose dependent role on setting heart rate. Collectively, this is compelling evidence for the involvement of RYR2 in pacemaking. One of the competing models for heart rate, the ‘two-clock hypothesis’ suggests that heart rate is driven by an ensemble of ion channels both in the plasma membrane and localized to the SR membrane of pacemaker cells in the SA node (Monfredi et al., 2013). A key aspect of this theory is that the ‘slow depolarization’ phase of pacemaker cell electrophysiology, which is the key determinant of heart rate, has a substantial contribution from RYR2 Ca2+ release driving depolarization through the NCX exchanger (Monfredi et al., 2013). As such, this model predicts that both RYR2 and the NCX channel play non-redundant roles in pacemaking (Monfredi et al., 2013). Our work here clearly demonstrates that RYR2 has a non-redundant role in heart rate, as decreases in channel levels diminish heart rate. Additional work, done concurrent to this thesis, has demonstrated that total deletion of NCX in the heart atria completely abolishes SA node pacemaking, and that hearts lacking atrial NCX must utilize a slow junctional escape rhythm mechanism to maintain heart beating (Groenke et al., 2013). This study clearly demonstrates that NCX plays a critical, non-redundant role in creating pacemaking behavior even in the context of   135 mostly normal heart function (Groenke et al., 2013). Between this study and the work presented in this thesis, there is substantial evidence that SR Ca2+ release and NCX dependent depolarization are key aspects of pacemaking, which provides key in vivo data to support the ‘two clock hypothesis’. There remains a great deal of work to be done to fully understand the role of RYR2 in pacemaking. The models described in this thesis, and in the vast majority of studies in the field, utilized a Mhy6 driven MerCreMer deletion system which generates robust recombination in pacemaker cells (Baruscotti et al., 2011), but also targets cardiomyoyctes throughout the heart (Sohal et al., 2001). Therefore these observations are whole organ effects. This is valuable information, however additional studies that utilize isolated sinoatrial pacemaker cells and more directly probe their electrophysiology would provide valuable insight into the role of RYR2 in this process. In addition, a study that utilized an atria-selective cre deletion system, such as the sarcolipin-cre system used in the NCX study (Groenke et al., 2013), would be an elegant approach to test whether the observed decreases in heart rate in my cRyr2KO and cRyr2Δ50 systems are directly caused by changes in sinoatrial pacemaking. Since loss of atrial contractility is not acutely fatal, this approach would also allow a test of whether sinoatrial pacemaking is lost in the complete absence of RYR2 in sinoatrial cells. I believe such a study program would be a valuable future direction of this work. Our results also indicate that loss of Ryr2 is arrhythmygenic. We observed that tachycardic episodes occurred in the cRyr2KO model but never in control animals. We did not perform the high-resolution ECG surveys or CPVT-inducing isoproterenol studies on the cRyr2Δ50 model needed to interrogate this process in the less dysfunctional model. This does create the possibility that the tachycardia we observed was a result of cardiac dysfunction instead of a direct signaling   136 effect of RYR2 reduction. However, given that some mutants have been observed which show diminished RYR2 activation and signaling and the generation of arrhythmias (Jiang et al., 2007; Thomas et al., 2004), I think this data supports the concept that reductions in RYR2 signalling can also disrupt cardiac rhythmicity. These results do not prove or suggest a mechanism, and further studies, again likely on isolated pacemaking cells would be needed to conclusively prove how a reduction in RYR2 may cause tachychardic arrhythmias.  How reductions in RYR2 signaling can cause tachycardia remains an intriguing question. I favor the model put forward by Jiang et al. (Jiang et al., 2007), which suggests that the arrhythmias may be caused by Ca2+ alternans. Alternans is a phenomena where Ca2+ signals, and therefore cardiac function, begin to alternate between big and small contractions and signals (Hüser et al., 2000; Picht et al., 2006; Surawicz and Fisch, 1992). It’s been proposed that in the presence of beta-adrenergic signalling these alternans may be sufficient to cause inappropriate NCX driven diastolic after-depolarizations (DADs) that can cause tachycardic arrhythmias (Hüser et al., 2000; Picht et al., 2006; Surawicz and Fisch, 1992).  The mechanism underlying alternans remains controversial, but it is generally elicited by alterations in SR Ca2+ release through SR Ca2+ load, RYR2 phosphorylation status, or metabolic changes (Hüser et al., 2000; Picht et al., 2006; Surawicz and Fisch, 1992). However, one model elicited Ca2+ alternans by using a gradual titration of tetracaine, an RYR2 inhibitor, to elicit the phenomena, suggesting that reducing RYR2 activation can potentially play a causal role (Diaz et al., 2002). In addition, when looking at the Ca2+ signaling in paced cRyr2Δ50 cardiomyocytes, we observed that Ca2+ waves showed much more gradual activation and deactivation, instead of the quick peaks seen in control cells. This is a phenomena not dissimilar to what is seen in alternans, and while it is far from conclusive, does suggest this model may warrant further investigation.    137 6.5   RYR2 and Metabolism The research presented in this thesis clearly establishes a role for RYR2 in promoting cardiac metabolism. With the cRyr2KO model we found that knocking out the Ryr2 gene resulted in decreased cardiac ATP levels, a general decrease in mitochondrial oxidative metabolism, and increased glycolysis. This is associated with reduced mitochondrial Ca2+ in unpaced cardiomyocytes, increased hypoxia factor signaling, and genetic reprogramming consistent with metabolic deficiencies. This provides compelling evidence that RYR2 acts to stimulate and maintain these processes and that loss of Ryr2 results in pseudohypoxia. However, the profound cardiac pathology observed in this model creates the possibility that much of this effect may be due to reduced cardiac function instead of a direct mechanistic link to RYR2. We therefore measured metabolism in the cRyr2Δ50 model, which displayed relatively well preserved cardiac function and a lack of obvious pathology. In this context we did not observe an overall decrease in cardiac ATP levels or general reductions in oxidative metabolism, but did see a specific decrease in glucose oxidation and increase in overall glycolysis. We further found that mitochondrial Ca2+ was diminished and partially uncoupled from cytoplasmic Ca2+ transients in cRyr2Δ50 paced cardiomyocytes.  In addition, we observed that PDH, the Ca2+ sensitive gatekeeper between glucose oxidation and glycolysis, was hyperphosphorylated and therefore less active in fasted cRyr2Δ50 hearts compared to fasted controls. This suggests a mechanism where RYR2 is required to drive mitochondrial Ca2+ uptake and stimulate the activating dephosphorylation of PDH, which then promotes increased glucose oxidation. Moreover, this research shows that RYR2 may play an important role in specifically driving glucose use as an energy substrate in hearts.   138 These findings are largely consistent with an emerging understanding of the link between cardiac Ca2+ signaling and heart metabolism. While in vitro work has long demonstrated that mitochondrial Ca2+ entry promotes oxidative metabolism (Denton, 2009; Glancy and Balaban, 2012), it is only recently, concurrent to the research contained in this thesis, that the genetics and molecular tools have become available to properly interrogate this system in vivo (Baughman et al., 2011; Drago et al., 2011; Kwong et al., 2015; Pan et al., 2013; Patron et al., 2013). These findings demonstrate that RYR2 is upstream of mitochondrial Ca2+ homeostasis, promoting glucose oxidation, and possibly stimulating general oxidative metabolism. Concurrent studies have shown the important of SR-mitochondrial Ca2+ by disrupting the signaling nanodomain and showing reductions in energy production in cardiac systems (Chen et al., 2012; Kohlhaas and Maack, 2010), and demonstrated that mitochondrial Ca2+ uptake and energy production occur on a beat-to-beat basis (Lu et al., 2013). Other very recent work deleting MCU, the main mitochondrial Ca2+ uptake mechanism, reveals that a complete loss of rapid mitochondrial Ca2+ uptake diminishes cardiac ATP production from rapid changes in demand (Kwong et al., 2015; Luongo et al., 2015; Wu et al., 2015), and that at least in certain contexts, causes a hyperphosphorylation and repression of PDH (Luongo et al., 2015; Pan et al., 2013). When taken together, this body of work establishes a model where RYR2 is upstream of mitochondrial Ca2+ and ATP production and highlights that this process is important to cardiac biology.  One of the interesting and unresolved questions raised by this work is whether RYR2 has a general role in promoting oxidative ATP metabolism or only a role in driving glucose oxidation. In the cRyr2KO study we saw significant decreases in glucose, lactate, and palmitate oxidation, as well as a general decrease in ATP levels. This would seem to indicate that ablation of RYR2 plays a general role in promoting mitochondrial ATP production by all oxidative substrates.   139 However, this model displayed significant heart dysfunction, and when we studied the milder cRyr2Δ50 model, we only observed decreased glucose oxidation. This creates the possibility that, physiologically speaking, RYR2 only has a role in promoting ATP production via glucose oxidation and that the general defects we saw in the cRYR2KO model were pathophysiology. That said, if the difference between these models is not due solely to pathology, then it is possible that Ca2+ and RYR2 may have a dose dependent role in mitochondrial ATP production. Perhaps a certain level of Ca2+ entry stimulates elements of the TCA cycle, ETC, and ATPase, to generally promote oxidation, but that glucose oxidation is more sensitive to changes in mitochondrial Ca2+ homeostasis and therefore more reliant on RYR2 to promote activation. This idea is somewhat born out by the in vitro and isolated mitochondrial studies which note that PDH has a higher sensitivity to Ca2+ than other TCA cycle Ca2+ sensitive dehydrogenases (Denton, 2009), as well as requiring a higher level of Ca2+ for full activation than the ETC elements and ATPase (Glancy et al., 2013).  PDH Ca2+ sensitivity is also independent of ATP levels, while other TCA enzymes become more sensitive to Ca2+ when cellular ATP levels are diminished (Denton, 2009), which may account for some of the phenotypic differences we observe between the cRyr2KO and cRyr2Δ50 models. The extra Ca2+ sensitivity of glucose oxidation may represent a regulatory mechanism by which ECC signalling can preferentially upregulate glucose utilization during periods of increased cardiac work (Kodde et al., 2007). Ultimately, our research does not provide data to answer this question and additional studies; perhaps targeted MCU knockout and haploinsufficiency models, which would allow modulation of mitochondrial Ca2+ uptake outside the context of heart failure (Kwong et al., 2015; Pan et al., 2013), would be needed to clearly establish whether mitochondrial Ca2+ has discrete dose dependent effects.   140 Another intriguing question raised by this research is why the effects of RYR2 ablation on PDH phosphorylation are context dependent. We observed that reduced RYR2 levels only resulted in hyperphosphorylated PDH complex when cRyr2Δ50 mice were fasted overnight. This finding is similar to what is seen in the constitutive MCU knockout study which also only observed PDH hyperphosphorylation and reduced enzyme activity in fasted mice (Pan et al., 2013). Since MCU knockout completely abrogates rapid Ca2+ uptake in mitochondria, it is not unreasonable to expect them to see a more profound metabolic phenotype than what we observe with our cRyr2Δ50 hearts (Pan et al., 2013).  Additionally, the heart specific MCU study did not observe any changes in PDH phosphorylation, although this study did not check this in the context of fasting or another metabolic stress (Kwong et al., 2015). This research suggests that PDH activation state is highly context dependent and may rely more on the PDK inhibitory arm of signaling than the proposed Ca2+ dependent activation via PDP1 dephosphorylation. I would speculate that under normal metabolic conditions, PDK activity is kept at a relatively low level in cardiac tissues, repressed by normoxia, ATP demand, and insulin signaling (Holness and Sugden, 2003).  In this context, low PDK activity leads to low levels of PDH phosphorylation and a low requirement for reactivating PDP activity, which may be satisfied by the constitutively active, but lower turnover PDP2 isoform (Huang et al., 1998). For Ca2+-dependent PDP1 dephosphorylation, and therefore RYR2 signaling, to become relevant to PDH activation there may be a requirement for a metabolic stress to activate PDK activity. While very much speculation at this point, this is not inconsistent with our observations since fasting should lead to PDK upregulation via reduced insulin signalling (Huang et al., 2002), and the reduced glucose oxidation we see in the working heart may involve hypoxic stress caused by the organ isolation process that cannot be compensated for in RYR2 ablated hearts. If this is the case, RYR2 and   141 PDP1 may act less to generally promote glucose oxidation but rather to de-repress PDH during metabolic stress caused by hypoxia or nutritional deficit. This may make this RYR2/PDP1/PDH signaling axis important during ischemia or increased workloads when glucose oxidation is preferentially upregulated despite a predicted increase in PDK signaling. Indeed, research using dicholoracetate or PDK4 knockout to repress PDK signaling during ischemia has found that de-repressing PDH activation significantly improves outcomes by better coupling glycolysis to glucose oxidation (McVeigh and Lopaschuk, 1990; Michelakis et al., 2002; Ussher et al., 2012). Further research into Ca2+ signaling and the context dependent regulation of PDH, as well as the relative contributions of PDP1 and PDP2 to heart metabolism will prove to be valuable to understanding how the heart utilizes glucose. The apparent disconnect between lactate and glucose oxidation in our cRyr2Δ50 model is potentially interesting. It has previously been shown that cardiac tissues will simultaneously take up lactate for oxidation in the TCA cycle while also exporting lactate generated as a product of glycolysis(Chatham et al., 2001). In addition, it has been found that cardiac mitochondria express MCT1 and matrix LDH1, suggesting that cytosolic lactate can be effectively taken up by mitochondria, converted to pyruvate, and utilized in the TCA cycle (Brooks et al., 1999). It has been proposed that lactate energy metabolism has been compartmentalized in the heart with glycolytically-derived lactate and exogenous lactate having distinguishable fates and different metabolic efficiencies (Brooks et al., 1999). This concept has been born out by the observation that exogenous lactate is consumed ~10-40% more efficiently by oxidative metabolism than exogenous pyruvate (Brooks et al., 1999; Molé et al., 1978), and that carbon from glyocolytically-derived pyruvate, but not exogenous lactate, can be found incorporated into other biomolecules (Chatham et al., 2001). It has even been proposed that glycolytically-derived   142 pyruvate is entirely converted to lactate to support glycolysis rates, and that any glucose derived carbons that are oxidized in the TCA cycle arrive at the mitochondria in the form of lactate(Brooks et al., 1999). Since we see decreased glucose oxidation rates, but no changes lactate oxidation rates, our cRyr2Δ50 results would support the theory that exogenous lactate and glycolytically-derived lactate are biochemically compartmentalized in the heart. Since it is thought that localized glycolysis may play a role in supporting ion homeostasis (Aasum et al., 1998; Xu et al., 1995), further research into this lactate compartmentalization in cRyr2Δ50 mice using stable-isotope working heart experiments may provide valuable insight into the phenomenon and the relationship between metabolism and cardiac function.  The research contained in this thesis, collectively suggests that RYR2 may act as a key integration point of cardiac function and metabolism. The data containd in this thesis clearly indicates that RYR2 is upstream of at least some aspects of ATP production in cardiomyocytes. Concurrent work by other groups examining the importance of mitochondrial Ca2+ on oxidative metabolism have additionally shown a clear linkage between SR Ca2+ release, mitochondrial Ca2+ homeostasis, and energy production (Chen et al., 2012; Kohlhaas and Maack, 2010; Kwong et al., 2015; Liu and O'Rourke, 2008; Lu et al., 2013; Pan et al., 2013). This suggests that RYR2 plays a critical role in coupling ECC directly to ATP production, and that this is part of the mechanism by which energy production is exquisitely matched to the metabolic demand of generating contractile force and maintaining the ionic gradients needed for ECC signaling. RYR2 gating is stimulated by binding ATP, making the channel sensitive to overall cellular energy load (Meissner et al., 1986).  In addition, other work has shown that RYR2 channel gating can be altered by oxidation and nitrosylation of various cysteine residues as a consequence of cell metabolism (Donoso et al., 2011; Oda et al., 2015). This suggests that   143 RYR2, and therefore ECC and cardiomyocyte contraction, are sensitive to and regulated by the metabolic status of the cell. This means that RYR2 may simultaneously regulate and be regulated by metabolism and may therefore represent a key integration point in the crosstalk between cellular function and energy production. As such, RYR2 signaling may be fundamental to cardiac biology in ways beyond its role in ECC coupling. Furthermore, since metabolism, contraction, and RYR2 function are often disrupted in heart failure, this function/metabolism crosstalk may prove to also be critical to cardiac pathology.  6.6   RYR2 and Cell Death The research presented in this thesis shows a substantial increase in cell death in the cRyr2KO model. This cell death is not associated with increased levels of cleaved caspase 3, or increased ER stress signaling, suggesting that this is not apoptotic cell death. We did, however, observe an increased amount of cleaved calpain 10 protein, at sizes consistent with an active protease. This is consistent with research conducted in the pancreatic beta cell which showed that pharmacological inhibition of RYR2 lead to atypical cell death that was abrogated on a calpain 10 deficient background. As such our data supports the model where RYR2 signalling is a pro-survival signal that represses activation of mitochondrial calpain 10 activation. While our data is consistent with calpain 10 activation, it critically does not prove this process is the sole driver of cell death or that calpain 10 activation is required for this process. While there is considerable evidence that calpain 10 is localizes to the mitochondria and shows Ca2+ sensitivity for activation (Arrington et al., 2006; Johnson, 2004; Ma et al., 2001), the exact mechanism for protease activation remain controversial (Arrington et al., 2006). Calpain 10 is considered an atypical calpain since it lacks the canonical ‘calpain-domain’ but still demonstrates   144 Ca2+ sensitive protease activity (Arrington et al., 2006). Our data, along with some previous studies (Johnson, 2004), would support that calpain 10 is generally repressed by Ca2+ signaling and becomes active when mitochondrial Ca2+ levels fall below a certain threshold. The exact molecular targets and the mechanism for how calpain 10 might mediate cell death also remain largely unknown. It is known that calpain 10 activation leads to DNA cleavage and laddering (Johnson, 2004), and indeed nuclear localized calpain 10 has been observed in certain models (Ma et al., 2001) but not others (Arrington et al., 2006), suggesting that calpain 10 entry to the nucleus may be involved in the process. The activation of other mitochondrial calpains are associated with opening of the MPTP (Gores et al., 1998; Polster et al., 2005), however calpain 10 over-expression studies have shown only small effects on this phenomenon, and I would suggest that these may have been experimental artifacts (Arrington et al., 2006). Additionally, calpain 1 and calpain 2 are activated by increased mitochondrial Ca2+ and we see no evidence of self-cleavage and activation in our model (Goll et al., 2003), suggesting that these proteases are likely not contributing to our phenotype. Clearly additional work is needed to uncover the exact mechanism of calpain 10 activation and to demonstrate what cell death effects are downstream of protease activation. Moreover, repeating these studies in a calpain 10 knockout model, with and without RYR2 ablation would allow for conclusive proof of calpain 10 involvement in Ryr2 deficient cell death and allow for further mechanistic insight. Another potentially interesting aspect of this study is the apparent reduction in SR/ER-stress signalling in the cRyr2KO model. Rather than see an increase in SR/ER-stress mediators of cell death, we actually observe a decrease; a phenomenon my supervisor has taken to calling ‘ER Zen’. This may be associated with an increase in SR Ca2+, since we have a reduction in RYR2, an ER Ca2+ release channel, although we lack direct SR Ca2+ measurements to support this.   145 Conversely, this decrease in ER-stress signaling may be a result of reduced RYR2 production. RYR2 is a very large protein (2.2 mDa/tetramer) and is highly expressed in cardiomyocytes (Lanner et al., 2010). This means that RYR2 probably exacts a considerably biosynthetic burden on the cell type. When insulin, the primary synthetic product of beta cells, is knocked out of pancreatic beta cells, the cells demonstrate ‘ER Zen’ and increased cell replication due to excess biosynthetic resources (Szabat et al., 2015). It is an interesting question of whether ablation of RYR2 production, a major protein product of cardiomyocytes, has similar effects in the heart.  6.7   RYR2 and Hypertrophy The data presented in this thesis suggests that loss of Ryr2 causes hypertrophy. Specifically we observed that Ryr2 deletion caused enlargement of the heart and an increase in heart weight, that was not associated with an increase in cytosolic Ca2+ in resting cardiomyocytes. We did not measure the molecular signaling system downstream of this apparent hypertrophy. This result is somewhat unexpected since previous models have implicated RYR2 signaling via the calcineurin/NFAT or ERK/MAPK pathways (Yamaguchi et al., 2011; Zou et al., 2011), and Ca2+ dependent hypertrophic signaling has generally been associated with increased cytosolic Ca2+ levels (Heineke and Molkentin, 2006). This creates an interesting situation worthy of further study: how does a heart with apparently reduced Ca2+ signaling drive hypertrophic gene programs? Our model would seem to suggest that there is a hypertrophic gene program that bypasses the apparent requirement for increased cystolic Ca2+ levels. This suggests that there may be a Ca2+ independent modality to this process, or perhaps, a substantial involvement of perinuclear IP3Rs which may be able to function as a Ca2+ microdomain promoting hypertrophy without increases in bulk cytosolic Ca2+ (Arantes et al., 2012; Guatimosim et al., 2008; Wu et al.,   146 2006). Alternately, our results may be confounded by the fact we measured Ca2+ in unpaced, isolated cRyr2KO cardiomyocytes which may have caused us to underestimate the true shape and size of cytosolic Ca2+ signals. Cardiomyocytes are never quiescent during normal biology and SR Ca2+ load is partially dependent on function, which makes measurements of unpaced cardiomyocytes somewhat limited. Furthermore, given that cytosolic Ca2+ levels are the main mechanism for terminating L-type VGCC Ca2+ signals, it is possible that these channels are much more active in the context of reduced RYR2 signaling (Peterson et al., 1999). Assessing the contribution of L-type VGCCs on cytosolic Ca2+ levels would require measuring cytosolic Ca2+ in paced cells. This maintains the possibility that cytosolic Ca2+ may not be decreased, or may even be elevated in the cRyr2KO model. Indeed, I have often wondered if basal cytosolic Ca2+ levels would be elevated in our model, since maintaining a near contraction threshold level of Ca2+ would provide a satisfying mechanism for how cRyr2KO hearts are able to maintain contractile function in the context of reduced SR Ca2+ release. Regardless, further research will be needed to establish the exact molecular mechanism for the hypertrophy observed in this study.  6.8   RYR2 and Cardiac Pathology A clear result of Ryr2 deletion was the induction of a rapid cardiac pathology that lead to death in cRyr2KO mice. This was associated with reduced cardiac function, slower heart rate, and arrhythmias. We also saw increased heart size that was associated with ventricular dilation and substantial fibrosis. Since our sole manipulation was the deletion of Ryr2, this data clearly reveals that loss of Ryr2 is disastrous for heart function and decreasing RYR2 signaling is able to recapitulate heart failure. This is strong evidence that the RYR2 dysfunction observed in several models of heart disease and aging (Assayag et al., 1998; Bidasee et al., 2001; Brillantes et al.,   147 1992; Crossman et al., 2011; Dincer et al., 2006; Kandilci et al., 2011; Matsui et al., 1995; Milnes, 2001; Naudin et al., 1991; Shao et al., 2007; 2012; Tellez et al., 2011; Wu et al., 2012; Yu et al., 1994) is indeed be pathogenic and can contribute to the development and progression of heart failure. An interesting issue of Ca2+ homeostasis in heart failure is whether reported RYR2 dysfunction or depleted SR Ca2+ load, which is seen in many models of heart failure, are drivers of pathophysiology. While some models of heart failure report reduced RYR2 levels, expression, and function (Bidasee et al., 2001; Brillantes et al., 1992; Crossman et al., 2011; Dincer et al., 2006; Matsui et al., 1995; Milnes, 2001; Naudin et al., 1991; Shao et al., 2007; 2012; Wu et al., 2012; Yu et al., 1994), other models do not report changes in the Ca2+ channel but instead a reduction in SR Ca2+ release caused by reduced SR Ca2+ load from increased NCX expression and reduced SERCA2 activity (Balke and Shorofsky, 1998; Bers, 2006; Gomez, 1997).  Our work reduces RYR2 levels and therefore signaling, and suggests that RYR2 dysfunction can be a driver of heart disease. However, our study does not directly measure SR Ca2+ load and cannot distinguish whether Ryr2 deletion also decreases SR Ca2+ stores. Therefore, further studies are necessary to tease out whether it is reduced SR Ca2+ release or SR store depletion that contribute to pathophysiology. Heart failure is a multifaceted condition associated with numerous defects. Besides the obvious reductions in cardiac contractility and function, heart failure is associated with morphological changes in the size and shape of the heart (McMurray and Pfeffer, 2005). In addition, heart failure is also associated with a decline in heart rate and incidences of arrhythmia (Fei et al., 1994; Ponikowski et al., 1997), as well as changes in cardiac metabolism and cell death (Stanley and Chandler, 2002). In our cRyr2KO model we observed decreased cardiac   148 function, but also alterations in heart rate, metabolism, and cell survival. Since we also observed that RYR2 stimulates oxidative metabolism and heart rate in the absence of heart failure, it is likely that RYR2 dysfunction can contribute to the lesions in oxidative metabolism and heart rate observed in heart disease. RYR2 may therefore be a key integration point of cardiac pathophysiology, where derangement of channel activity can simultaneously disrupt multiple downstream aspects of cardiomyocyte biology. Moreover, the fact that RYR2 is potentially regulated by both cardiac function (Lanner et al., 2010; Meissner, 1994) and metabolism (Donoso et al., 2011; Meissner et al., 1986), means the channel may act as an important link between metabolic and contractile dysfunction, and can perhaps facilitates how deficiencies in one process progress to affect the other. Collectively my thesis work suggests that RYR2 dysfunction is potentially involved in numerous aspects of heart failure, or, at the very least, that Ryr2 loss can recapitulate a complex heart failure phenotype. A particularly interesting aspect of this data is the link between RYR2 and metabolism. We observed that Ryr2 deletion resulted in heart failure as well as a general reduction in available ATP and oxidative energy metabolism. We further saw, using the cRyrΔ50 model, that a stable reduction in RYR2 specifically reduced glucose oxidation and increased glycolysis rates. This suggests RYR2 may play a role in promoting oxidative metabolism generally and to specifically couple glucose oxidation to glycolysis through activating, or de-repressing, the PDH complex. Oxidative energy metabolism is generally decreased in heart failure (Neubauer, 2007; Stanley and Chandler, 2002), and a variety of studies have suggested that energy substrate selection is altered in various models of heart disease (Neubauer, 2007; Stanley and Chandler, 2002). Two particularly interesting findings are that glucose oxidation is virtually abolished in experimental models of diabetic cardiomyopathy (Lopaschuk, 2002) and that glucose oxidation and glycolysis   149 frequently become uncoupled in heart failure (Stanley and Chandler, 2002). RYR2 dysfunction has been demonstrated in models of both diabetic cardiomyopathy (Bidasee et al., 2001; Crossman et al., 2011; Dincer et al., 2006; Shao et al., 2007; 2012; Wu et al., 2012; Yu et al., 1994), and pressure-overload heart failure (Brillantes et al., 1992; Matsui et al., 1995; Milnes, 2001; Naudin et al., 1991), which in the context of the work presented here, suggests that RYR2 may play an underappreciated role in contributing to these metabolic effects. Given that promoting glucose oxidation in failing or ischemic hearts has been shown to ameliorate cardiac injury and pathophysiology (McVeigh and Lopaschuk, 1990; Michelakis et al., 2002; Ussher et al., 2012), understanding the role of RYR2 in this process may be clinically important and may provide a new avenue for therapeutic intervention. Given that this research establishes a role for RYR2 dysfunction in contributing to multiple aspects of cardiac biology and heart failure, this thesis implicates RYR2 as an attractive potential therapeutic target for heart failure. Since RYR2 is upstream of cardiac contraction, heart rate, and energy metabolism, modifying RYR2 activity may allow for an integrated approach that can potentially benefit numerous facets of heart disease. Indeed, some groups are hoping to modify RYR2 function as a potential way to lower heart rate and abolish arrhythmias (Dulhunty et al., 2011; Hwang et al., 2011; Mackrill, 2010; Yamamoto et al., 2008). Flecainide, a drug used to acutely treat arrhythmias, inhibits both SCN5 Na+ channels and RYR2 (Hwang et al., 2011). However, any modification of RYR2 behavior has to be exquisitely careful. Increased RYR2 activity has been linked to fatal catecholaminergic arrhythmias (Blayney and Lai, 2009), hypertrophy (Yamaguchi et al., 2011), and heart failure (Yamaguchi et al., 2007), showing that excessive RYR2 activity is dangerous and pathogenic. This thesis showcases that RYR2 is simultaneously involved in many core biological processes and that reducing RYR2 expression   150 and function is also pathogenic. Collectively this argues that RYR2 signaling is a ‘goldilocks’ phenomenon and that any pharmacological intervention of the channel has to be careful to maintain an optimal range of signaling. Moreover, this research also argues that any potential therapeutic modification of RYR2 must be cognizant of the reality that the channel regulates many cardiac processes and that targeting RYR2 to change one aspect of heart function will have off-target consequences. Overall, RYR2, while an attractive candidate, may ultimately be too central and too potentially dangerous to be a viable therapeutic target for heart disease in most circumstances.  6.9   Overall Significance This thesis demonstrates that RYR2 has a varied and central role in cardiac biology. It provides new, in vivo evidence that RYR2 is necessary for cardiac function in adult animals and provides evidence that loss of Ryr2 and diminished RYR2 signaling is pathogenic. The most significant findings, however, are that RYR2 is plays a fundamental role in cardiac pacemaking and that RYR2 signaling promotes oxidative metabolism. This work showing that RYR2 deletion or knockdown reduces heart rate, along with work revealing that NCX is critical for SA node pacemaking (Groenke et al., 2013), establishes profound evidence to support the ‘two-clock hypothesis’ of heart rate and an ensemble mechanism of cardiac pacemaking (Monfredi et al., 2013). This work, along with a variety of congruent studies (Chen et al., 2012; Kohlhaas and Maack, 2010; Kwong et al., 2015; Liu and O'Rourke, 2008; Lu et al., 2013; Pan et al., 2013), also establishes a model where RYR2 Ca2+ release drives mitochondrial Ca2+ uptake to promote oxidative ATP production. Critically, this work also demonstrates that RYR2 has a role in promoting glucose oxidation by driving the full activation of the PDH complex, thus linking   151 glycolysis with the full oxidation of this energy substrate. This provides a novel mechanism for the heart to directly couple glucose oxidation to cardiac function and potentially de-repress glucose use during metabolic or hypoxic stress. This thesis also provides intriguing evidence that RYR2 is involved in cardiac hypertrophy and cell death via a calpain 10 dependent mechanism. Collectively this suggests that RYR2 sits at a nexus of cardiac processes and plays an important role coupling general cell biology to cardiac function. This, I think, creates a model of RYR2 where it functions as a key integration point of cardiomyocyte biology. RYR2 has important roles in ECC signaling meaning that RYR2 is a critical driver of cell contraction and overall cardiac function. The same RYR2 signals also have an apparent role in stimulating cell metabolism and energy production as well as cardiac hypertrophy (Yamaguchi et al., 2011; Zou et al., 2011). Therefore, RYR2 can couple processes that support heart function directly to cardiomyocyte contraction, thus providing an exquisitely efficient method to ensure the metabolic and biosynthetic requirements of cardiomyocyte contraction are constantly satisfied. In addition, RYR2 channel gating is regulated by a variety of molecules that allow it to respond to changing conditions. RYR2 activity is modified by a variety of Ca2+ sensor molecules (Bers, 2004), is downstream of beta adrenergic signaling (Meissner, 2002), and is able to sense SR and cytosolic Ca2+ directly (Chen et al., 2014), meaning that RYR2 activity is modified by cardiac function.  RYR2 is also able to sense metabolism through direct interaction with ATP (Meissner et al., 1986), oxidation and nitrosylation as a byproduct of energy metabolism (Donoso et al., 2011), and indirectly via the effects of SERCA activity on SR Ca2+ load. This means that RYR2 both regulates and is regulated by cardiac function and metabolism such that the channel potentially bridges these two facets of cardiac biology and provides a mechanism for co-regulation. Collectively, I think this favors a model where RYR2   152 functions not so much as a discrete ECC signaling molecule, but rather as a signaling nexus that manages several key aspects of cardiac function and directly couples cardiomyocyte contraction to the general cell biology needed to support it.    153 Bibliography Aasum, E., Lathrop, D.A., Henden, T., Sundset, R., and Larsen, T.S. (1998). The role of glycolysis in myocardial calcium control. J. Mol. Cell. Cardiol. 30, 1703–1712. Albu, R.F., Chan, G.T., Zhu, M., Wong, E.T.C., Taghizadeh, F., Hu, X., Mehran, A.E., Johnson, J.D., Gsponer, J., and Mayor, T. (2015). A feature analysis of lower solubility proteins in three eukaryotic systems. J Proteomics 118, 21–38. Allard, M.F., Schonekess, B.O., Henning, S.L., English, D.R., and Lopaschuk, G.D. (1994). Contribution of oxidative metabolism and glycolysis to ATP production in hypertrophied hearts. Am. J. Physiol. 267, H742–H750. 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