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Dynamic distribution of ryanodine receptors in cardiac muscles Asghari, Parisa 2014

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 Dynamic Distribution of Ryanodine Receptors in Cardiac Muscles   by Parisa Asghari     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)   September 2014  © Parisa Asghari, 2014   ii Abstract The focus of this thesis is to address the location and distribution of the type 2 Ryanodine Receptor (RyR2) in mammalian cardiac myocytes with respect to their function. These integral membrane proteins function as Ca2+-activated Ca2+ ion channels as well as a scaffold for a large number of signaling molecules that modulate the release of Ca2+ through the channel. The relative position of the RyR2 tetramers is therefore a critical determinant of their function.  To study this question, I have used a combination of immunofluorescence microscopy, transmission electron microscopy, and tomography to map the position of the tetramers in whole cells and in cell sections and have used tissue obtained from both rat and human hearts. Biochemical and physiological techniques were used to correlate structure with function. I have found that RyR2s are located only in three regions: in couplons on the surface, transverse tubules and on most of the axial tubules. In all regions, most but not all of the RyR2s colocalize with the voltage-gated Ca2+ channel (Cav1.2), suggesting that they play a role in excitation-contraction coupling. Some RyR2 are colocalized with cavelin-3 and not with Cav1.2 and hypothesized that these ‘extra-couplonic’ RyR2 might be regulated by the multitude of signaling molecules associated with caveolin-3 to modulate Ca2+ release. Dual-tilt electron tomography produced en face views of both rat and human dyads, enabling a direct examination of RyR2 arrangement. Both species showed that tetramer packing was non-uniform containing a mix of checkerboard and side-by-side arrangements as well as isolated tetramers. Finally, I showed that the tetramers’ arrangement depended on the Mg2+ concentration and on their phosphorylation status; in low Mg2+ and after phosphorylation RyR2s were positioned in largely checkerboard arrangements while in response to high Mg2+ the tetramers were positioned largely side by side. These tetramer arrangements: side by side, mixed and checkerboard were associated   iii with progressively increasing spark frequencies. The correlation between tetramer arrangement and spark frequency suggests that tetramer rearrangement may be another mechanism whereby physiological processes operate and provides potential new mechanisms by which the activity of RYR2, the dyad and cardiac contractility may be regulated.      iv Preface This thesis is composed of work that has been published in peer-reviewed journals. Some part of the Introduction (Chapter 1) and Conclusion (Chapter 5) were adapted from a published review article: Scriven DRL, Asghari P, Moore EDW. Microarchitecture of the Dyad. Cardiovasc Res. 2013;98:169–176. Chapters 2, 3 and 4 are published articles and presented in their original forms.   Chapter 2: Asghari, P., Schulson, M., Scriven, D.R., Martens, G., and Moore, E.D. (2009). Axial tubules of rat ventricular myocytes form multiple junctions with the sarcoplasmic reticulum. Biophys J 96, 4651-4660.  P. Asghari was responsible for designing and performing experiments, data analysis, producing figures, tables and writing of the manuscript in partnership with E.D. Moore. M. Schulson was involved in Figure 2.8 data acquisition and analyses. G. Martens helped with tissue preparation and electron tomography data acquisition. D. Scriven provided guidance and was involved in performing and analyzing immunefluorescence data in Figure 2.8 and writing the paper. Chapter 3: Asghari, P., Scriven, D.R., Hoskins, J., Fameli, N., van Breemen, C., and Moore, E.D. (2012). The structure and functioning of the couplon in the mammalian cardiomyocyte. Protoplasma 249 Suppl 1, 31-38. Reproduced with permission. P. Asghari was responsible for designing and performing experiments, data analysis, producing figures and writing of the manuscript in partnership with E.D. Moore. Authors, J. Hoskins, N. Fameli, C. van Breemen were involved in developing tetramer-fitting program used in Figure 3.3. D. Scriven provided guidance and was involved in analyzing the data and writing the manuscript.    v Chapter 4: Asghari, P., Scriven, D.R., Sanatani, S., Gandhi, S.K., Campbell, A.I., and Moore, E.D. (2014). Nonuniform and variable arrangements of ryanodine receptors within mammalian ventricular couplons. Circ Res 115, 252-262. P. Asghari performed and designed all the experimental procedures, including electron tomography, Western blots, Ca2+ spark collection, tetramer fitting, data analysis and also producing figures and tables. Human ethics approval (UBC REB # is H11-03319), patient contact and consent, and surgical samples were provided by S. Sanatani, S.K. Gandhi, A.I. Campbell. Authors: P. Asghari, D. Scriven and E.D. Moore wrote the paper and analyzed the data. D. Scriven wrote the tetramer-fitting program.    vi Table of Contents 	   Abstract .............................................................................................................................. ii	  Preface ............................................................................................................................... iv	  Table of Contents ............................................................................................................. vi	  List of Tables ...................................................................................................................... x	  List of Figures ................................................................................................................... xi	  List of Abbreviations ..................................................................................................... xiii	  Acknowledgements ........................................................................................................ xiv	  Dedications........................................................................................................................ xv	  1.	   Introduction ......................................................................................................................... 1	  1.1	   Excitation-contraction coupling ......................................................................... 1	  1.2	   Myocyte architecture ......................................................................................... 4	  1.2.1	   Ryanodine receptors .................................................................................... 7	   The macromolecular complex of RyR within the SR .......................... 8	  1.2.2	   L-type calcium channel ............................................................................. 10	  1.2.3	   The sodium-calcium exchanger ................................................................ 11	  1.2.4	   Caveolin 3 ................................................................................................. 12	  1.2.5	   Junctophilin ............................................................................................... 13	  1.3	   Ryanodine receptor’s structure and function ................................................... 15	  1.3.1	   Ultrastructure of the RyR .......................................................................... 16	   Movements associated with channel gating and coupled gating ....... 16	  1.3.2	   RyR arrays ................................................................................................ 19	    vii 1.3.3	   Regulation of RyR2 .................................................................................. 20	   Calcium, magnesium and ATP .......................................................... 21	   FKBP 12 ............................................................................................ 23	   Calmodulin ......................................................................................... 24	   Phosphorylation ................................................................................. 25	   RyR2 phosphorylation and cardiac disease ................................ 27	   Mechanisms underlying different effects of PKA and CaMKII on RyR2 gating…….. .................................................................................................... 29	  1.4	   Research hypotheses and aims ......................................................................... 31	  1.4.1	   Chapter 2 ................................................................................................... 31	  1.4.2	   Chapter 3 ................................................................................................... 31	  1.4.3	   Chapter 4 ................................................................................................... 31	  2.	   Axial tubules of rat ventricular myocytes form multiple junctions with the SR .............. 32	  2.1	   Synopsis ........................................................................................................... 32	  2.2	   Introduction ...................................................................................................... 32	  2.3	   Methods............................................................................................................ 34	  2.3.1	   EM tissue processing ................................................................................ 35	  2.3.2	   Preparing sections for single-axis tomography ......................................... 36	  2.3.3	   Image acquisition and tomography ........................................................... 36	  2.3.4	   Fluorescence microscopy .......................................................................... 37	  2.4	   Results .............................................................................................................. 37	  2.5	   Discussion ........................................................................................................ 52	  3.	   The structure and functioning of the couplon in the mammalian cardiomyocyte ............. 57	    viii 3.1	   Introduction ...................................................................................................... 57	  3.2	   Hypotheses ....................................................................................................... 61	  3.2.1	   Couplons are diverse and some are variable amplifiers of ECC .............. 61	  3.2.2	   The RyR2 within a couplon form a disordered array ............................... 64	  4.	   Non-uniform and variable arrangements of ryanodine receptors within mammalian ventricular couplons .................................................................................................................. 71	  4.1	   Introduction ...................................................................................................... 71	  4.2	   Materials and methods ..................................................................................... 72	  4.2.1	   Experimental protocol ............................................................................... 73	  4.2.2	   Identification and placement of ryanodine receptors ................................ 73	  4.2.3	   Statistical analyses .................................................................................... 74	  4.2.4	   Supplemental method ................................................................................ 74	   Calcium sparks in permeabilized myocytes ....................................... 75	   Phosphorylation ................................................................................. 76	   Confirmation of RYR2 phosphorylation ........................................... 77	   In situ fixation .................................................................................... 78	   Preparation for tomography ............................................................... 78	   Image acquisition and tomography .................................................... 78	  4.3	   Results .............................................................................................................. 79	  4.4	   Discussion ...................................................................................................... 105	  5.	   General discussion .......................................................................................................... 111	  5.1	   Overview ........................................................................................................ 111	  5.2	   Merging the finding of this thesis work and its significance to the field ....... 112	    ix 5.2.1	   Cellular architecture and calcium release units ...................................... 112	  5.2.2	   Dyadic clefts are not filled with RyR2 ................................................... 117	  5.2.3	   RyR2 do not occur in regular arrays ....................................................... 119	  5.2.4	   Non-uniform arrangements ..................................................................... 121	   Cytoplasmic magnesium ........................................................... 122	   The effect of phosphorylation on channel distribution ............. 123	  5.2.5	   Channel movement ................................................................................. 128	  5.3	   Significance .................................................................................................... 128	  Bibliography ...................................................................................................................129	     x List of Tables Table 4.1    Tetramer arrangement ...........................................................................................................................88	  Table 4.2   Nearest neighbor centre-to-centre distances (nm) ................................................................................89	        xi List of Figures Figure 1.1 A Schematic diagram of the classical view of the couplons in ventricular myocytes. ..........................4	  Figure 1.2 Transmission electron micrographs of rat ventricular myocytes fixed in situ displaying junctions with various morphologies.. .........................................................................................................................................7	  Figure 1.3 CryoEM map of RyR2 tetramer as viewed from the myoplasm (A), and from the side (B). ............18	  Figure 1.4 Modulators of RyR2. ................................................................................................................................30	  Figure 2.1 Transmission electron micrographs of junctions in rat ventricle. .......................................................39	  Figure 2.2 Longitudinal sections of txial tubules and their junctions.  ..................................................................40	  Figure 2.3 Frequency histograms. .............................................................................................................................43	  Figure 2.4 Cross sections of axial tubules and their junctions. ...............................................................................44	  Figure 2.5 Atrial RyR and corbular SR.  ..................................................................................................................45	  Figure 2.6 Tomography of a single longitudinal junction.  .....................................................................................46	  Figure 2.7 Tomography of a single longitudinal junction and the transverse tubule to which it is connected. .48	  Figure 2.8 Short and long stretches of axial RyR and Cav1.2. ...............................................................................51	  Figure 3.1 Transmission electron micrographs of adult rat ventricular myocyte. ...............................................59	  Figure 3.2 Ultrastructure of a myocyte and a couplon. ...........................................................................................68	  Figure 3.3 Tomogram of individual couplon from adult rat ventricular myocyte. ..............................................70	  Figure 4.1 RYR2 distribution in a rat cardiomyocyte fixed in situ. .......................................................................82	  Figure 4.2 Two different sets of orthogonal views of the tomogram presented in Figure 4.1. ............................85	  Figure 4.3 .....................................................................................................................................................................86	  Figure 4.4 Displays histograms of the tetramer’s nearest neighbor distances for myocytes fixed in situ. .........87	  Figure 4.5  RYR2 distribution in a human left ventricular myocyte. ....................................................................92	  Figure 4.6   Ca2+ Sparks frequency and SR content. ...............................................................................................94	  Figure 4.7 RYR2 distribution in a permeabilized rat cardiomyocyte with a free Mg2+ concentration of 0.1 mM. ..............................................................................................................................................................................95	  Figure 4.8 Additional images of the tomograms for cells that were permeabilized and exposed to 0.1 mM Mg2+. .............................................................................................................................................................................96	    xii Figure 4.9 Additional images of the tomograms for cells that were permeabilized and exposed to 4 mM Mg2+........................................................................................................................................................................................97	  Figure 4.10 Additional images of the tomograms for cells that were permeabilized and exposed to cAMP with phosphodiesterase and phosphatase inhibitors. .......................................................................................................99	  Figure 4.11 RYR2 distribution in a permeabilized rat cardiomyocyte with a free Mg2+ concentration of 4.0 mM. Scale bars are 30 nm. .......................................................................................................................................101	  Figure 4.12 Western blots. ........................................................................................................................................103	  Figure 4.13 RYR2 distribution in a permeabilized rat cardiomyocyte dyad after phosphorylation Scale bars are 30 nm. ..................................................................................................................................................................104	  Figure 5.1 A schematic picture of my view of RyR2 distribution within dyad and extra couplonic RyR2. .....114	  Figure 5.2 3D Electron tomography reveals the nano-scale 3-D structure of dyads in rat ventricular myocytes.....................................................................................................................................................................................119	  Figure 5.3 New and old view of dyad. .....................................................................................................................121	  Figure 5.4 Phosphorylation of ser-2814 in response to increase of endogenous CaMKII in rat cardiac myocytes. ....................................................................................................................................................................126	  Figure 5.5 RyR2 distribution in a permeabilized rat cardiomyocyte with activated endogenous CaMKII. ...127	      xiii List of Abbreviations βAR                β 2 Adrenergic Receptor 2D                   Two-Dimensional 3D                   Three-Dimentional CaM                Calmodulin CaMKII          Calmodulin Kinase II cAMP             Cyclic AMP Cav-1              Caveolin-1 Cav-2              Caveolin-2 Cav-3              Caveolin-3 Cav1.1             Skeletal L-type Calcium Channel Cav1.2             Cardiac L-type Calcium Channel CICR              Calcium Induced Calcium Release CPVT             Catecholaminergic Polymorphic Ventricular Tachycardia CSQ               Calsequestrin CSQ1             Skeletal Calsequestrin CSQ2             Cardiac Calsequestrin cSR                Corbular Sarcoplasmic Reticulum DADs             Delayed After Depolarizations ECC                Excitation Contraction Coupling EM                  Electron Microscopy HF                   Heart Failure INa                   Current through the voltage-gated Na+ Channel INCX                 Current through the Na+/Ca2+ Exchanger JPH                 Junctophilin JPH1               Brain Junctophilin JPH2               Cardiac Junctophilin JPH3               Skeletal Junctophilin jSR                  Junctional Sarcoplasmic Reticulum KO                  Knock-Out MI                   Post-Myocardial Ischemia MORN            Membrane On Receptor Nexus MY                  Myoplasmic Area NCX                Na+/Ca2+ Exchanger NND                Nearest-Neighbor center-to-center Distances RyR1               Type 1 Ryanodine Receptor (Skeletal) RyR2               Type 2 Ryanodine Receptor (Cardiac) RyR3               Type 3 Ryanodine Receptor SERCA           Sarcoplasmic Reticulum Ca2+ ATPase SOICR            Store Overload-Induced Calcium Release TATS             Transverse-Axial Tubule System TEM               Transmission Electron Microscopy TM                 Transmembrane Domain    xiv Acknowledgements I would like to take this opportunity to thank the many people who have helped me throughout the course of this degree. Without them, this simply would not have been possible.  My sincere gratitude must first go to my supervisor, Dr. Edwin Moore. His constant support, understanding, and generosity has kept me going through the rough patches, and his tremendous teaching ability has been invaluable.  Second, I would like to thank the other members of my supervisory committee Dr. Wayne Vogl, Dr. Patrick Nahirney and Dr. Filip Van Petegam for the time and energy they have committed, and the support they have given to this project.  I must extend many thanks to my fellow lab members, who have committed valuable time and effort in helping me finish this project. I must especially thank Dr. David Scriven, for enlarging my vision of science and providing coherent answers to my questions. His intellect and technical skills have helped me tremendously along my path, but most of all David was like a kind brother to me.  Finally, I would like to thank the most important people in my life, my family, my mother, husband, siblings and especially my son. Without their unconditional love and support I would not have been able to make it to this milestone; a PhD. My particular thanks also go to Dr. Sally Osborne, who has taught me so much. She has been an inspiration to me, but most of all, an extraordinary friend.     xv Dedications  This thesis is dedicated to Kevan and Darius. Their love, support, patience and understanding have lightened my spirit, enabling me to finish this study and this thesis.    1 1. Introduction It is a defining principle of cellular architecture that second messenger networks are spatially confined. Spatial confinement creates local domains for signal generation and termination that physically incorporate the key targets that will be regulated. Multiple signals can be then sent in parallel to many targets with slight or no cross talk between them, despite the fact the domains are created from the same basic components. Spatial confinement also ensures that the information is encoded. Since faulty signaling networks are responsible for a host of diseases, it is not surprising that uncovering their domains, and discovering how they are organized in both health and disease, has become a critical goal for cell physiologists.  Excitation-contraction coupling (ECC), regulated by the second messenger Ca2+, is an excellent model of how information can be spatially encoded. Once seen only from the perspective of the whole cell, the Ca2+ transient is now viewed as the macroscopic sum of thousands of spatially confined release and re-uptake events; an organization offering a rich environment for regulation that has only begun to be explored.   1.1 Excitation-contraction coupling ECC in ventricular myocytes initiates through an influx of Ca2+ through voltage-gated Ca2+ channels (Cav1.2), also known as dihydropyridine receptors, on the sarcolemma. The Ca2+ triggers a considerable amount of Ca2+ release into the myoplasm through cardiac ryanodine receptors situated in the membrane of the junctional sarcoplasmic reticulum (jSR), and this process of ECC is called Ca2+-induced Ca2+ release (CICR) (Fabiato, 1983, 1985). The structural component shaped by apposition (10-15nm) of the sarcolemma and jSR is called a dyad (Carl et al., 1995). RyR2 in jSR and Cav1.2 in the adjacent sarcolemma form a functional element within   2 the dyad known as a couplon (Figure 1.1A). Myocytes smoothly grade their free myoplasmic Ca2+ concentrations and the force of contraction as a function of membrane voltage (Em), and this is a hallmark feature distinguishing CICR from the voltage-gated mechanism of ECC in skeletal muscle (Beuckelmann and Wier, 1988). In skeletal muscles, type1 ryanodine receptors (RyR1) in jSR and skeletal voltage-gated Ca2+ channels (Cav1.1) in the sarcolemma interact directly, through electro-mechanical coupling.   Although CICR is a process with positive feedback, an open RyR2 does not activate all of the other channels leading to an explosive all-or-none event. This puzzle, the Ca2+ paradox, was explained theoretically by placing RyR2 into groups of functionally and physically separated clusters that were separately controlled (Stern, 1992). This was followed in rapid succession by experimental support. First came a series of observations describing Ca2+ ‘sparks’, which are the result of random openings of a small cluster of RyR2, which do not proceed to all-or-none Ca2+ transients (Cannell et al., 1994, 1995; Cheng et al., 1995; Chopra et al., 2009). These were spatially separated Ca2+ release events that did not activate Ca2+ release from other dyads. Following this, immunofluorescence and electron microscopy analyses characterized clusters of Cav1.2 in the sarcolemma positioned facing clusters of RyR2 in the jSR, within a distance of 12-­‐15 nm distance (Carl et al., 1995; Sun et al., 1995). The structural analyses and the Ca2+ sparks offered the basis for local control theory, which states that because of the physical separation between junctions and the high local Ca2+ required to activate RyR2, Ca2+ release from one junction does not affect the others. The central event in ECC is a Ca2+ spark, and a Ca2+ transient is the macroscopic summation of thousands of sparks. This allows the force of contraction to be a result of the number of sparks that are initiated, corresponding to the number of couplons that are engaged. As these theories were developed, a dyad was thought to cover a single large couplon,   3 and the RyR2s within a couplon could all contribute equally to a spark. However, the results of my research summarized in this dissertation, and the work of others, indicate that this is unlikely to be true (Chapter 4).  Cardiac muscle relaxation is the result of the removal of cytosolic Ca2+ by the sarcoplasmic reticulum Ca2+ ATPase (SERCA) pump and the Na+-Ca2+ exchanger (NCX). SERCA function is regulated by two small molecular weight proteins: phospholamban and sarcolipin (Vangheluwe et al., 2006). NCX is a secondary active transporter, importing three Na+ ions in exchange for one Ca2+ ion across the sarcolemma.        4   Figure 1.1 Schematic diagram of the classical view of the couplons in ventricular myocytes.  jSR was thought to be located almost exclusively along the surface membrane and the t-tubules, running along the Z-lines, and this is where the majority of RyRs and Cav1.2 form dyads. (A) The classical view of RyR2 distribution within the cell. (B) Couplons are shown in outline (light patches). Pointed region in (A) rotated 90°. The distance between couplons is about 300nm.    1.2 Myocyte architecture The classical view of myocyte structure came largely from the work of Franzini-Armstrong, and from Peachey (Franzini-Armstrong and Peachey, 1981) and Page (Page, 1978) among many others (Figure 1.1). In the ventricles and the atria of larger mammalian species, with a well-developed transverse-axial tubule system (TATS) (Kuwajima et al.), dyads are found largely on the cell surface and around the Z-lines (Dibb et al., 2009; Richards et al., 2011).   ~300	  nm     T-­‐tubule Rotate	  90°  T-tubule  Sarcolemma jSR  cSR  RyR Cav1.2 Axial-tubule   5 Corbular SR (cSR) are structurally specialized domains extending from network SR and form sacs of SR decorated with RyR2 that are located along the Z-line. cSR is found in myocytes without t-tubules and CSQ is present in the lumen. It was thought that cSRs contain up to 40% of a ventricular myocyte’s RyR2 (Jorgensen et al., 1993) (Figure 1.1 A). In response to Ca2+ diffusion from sarcolemmal dyads, cSRs might release Ca2+ and function as a secondary amplification system.  jSR appears as a slim sac that is flattened next to the sarcolemma, but it can embrace a number of configurations around the tubular system, forming dyads, triads, and more complex geometries, including junctions that cross an entire sarcomere (Figure 1.1 a) (Kirk et al., 2003). The width of the dyadic cleft (the distance between the jSR and the sarcolemma, which is 12-15 nm) is critical for effective CICR and for Ca2+ dependent inactivation of Cav1.2.  The Ca2+ released from RyR2 inactivates the adjacent Cav1.2 to speed the rate of repolarization (Lee et al., 1985; Yue et al., 1990). RyR2s span most of the width of the dyadic cleft, occupying the surface of the jSR facing the sarcolemma. They also have been known as the foot processes. Their position and characteristic shape make them one of the few proteins that can be positively identified in transmission electron microscopy (TEM). The luminal side of the jSR contains other proteins, seen in TEM as electron dense material, which has been identified in part as the Ca2+-binding protein calsequestrin (CSQ) (Sun et al., 1995). Examples of various junction morphologies are presented in Figure 1.2. The organization of the TATS in rat cardiac myocytes is completed between 10 to 12 days of the developmental process (Ziman et al., 2010). Early examination using electron microscopy revealed that very few L-type Ca2+ channels appear across from the RyR early in development (Franzini-Armstrong et al., 2005). Examination of developing rabbit myocardium showed that   6 NCX co-localized with RyR2 in the periphery (Dan et al., 2007). With growth, NCX expression weakens (Franzini-Armstrong et al., 2005) while the density and number of RyR2 clusters co-localized with NCX decrease (Dan et al., 2007). At the same time there is an increased co-localization of Cav1.2 with RyR2 (Sedarat et al., 2000) constructing the mature form of CICR, where ICa triggers Ca2+ release from RyR2.  Junctophilin (JPH), which anchors the sarcolemma to the SR might be the first molecule that appears in the junction, then CSQ appears in the luminal side of the SR beside junctin and triadin (Franzini-Armstrong et al., 2005). It has been suggested that in rat ventricular cardiomyocytes, the TATS is formed in two ways: tubes enter into the myocytes along the Z-lines, while small pouches of membrane containing Cav1.2 that form within the myocyte interior join the developing transverse tubules (Di Maio et al., 2007).      7  Figure 1.2 Transmission electron micrographs of rat ventricular myocytes fixed in situ displaying junctions with various morphologies. (A) Junction encircles t-tubule; (B) sarcolemmal junction; (C) dyad; (D) triad; (E) surface junction at intercalated disc. RyR2, double arrow; CSQ, arrow; ec, endothelial cell; is, interstitial space; jSR, junctional sarcoplasmic reticulum; t, t-tubule; m, mitochondria; z, z-line. Insets are two-fold magnifications of the boxed regions. Scale bars = 50 nm.   1.2.1 Ryanodine receptors Ryanodine receptors are part of the large family of homotetrameric Ca2+-activated Ca2+ release channels. There are three tissue specific mammalian isoforms (RyR1-3). RyR1 is the dominant isoform in skeletal muscle, and RyR2 is the most abundant isoform expressed in cardiac muscles (Fill et al., 2000). RyR1 and RyR2 are essential for muscle contraction, but the role of RyR3 is not clear. RyR3 has been suggested to play a role in learning and memory and also has sr# t#mA#sr#t#zmD# E#mz#sr#mzt#C#zB#is# ec#  8 been reported to be present in the early development of skeletal muscle (Balschun et al., 1999). RyR2 is also expressed at small levels in adrenal glands, kidney, stomach, lungs, thymus and ovaries (Nakanishi et al., 1992; Sharp et al., 1993) and in high levels in Purkinje cells of cerebral cortex and cerebellum (Giannini et al., 1995). Cerebral RyR2 has been shown to have a fundamental role in memory processing (Galeotti et al., 2008).  The three RyR isoforms show a considerable homology in structure and function. The transmembrane domain (TM) is found in the C-terminal ∼800 residues, while the N-terminal and central domains with ∼4100 amino acids form the cytosolic portion (Fill et al., 2000). Cryo-electron microscopy of RyR1 demonstrates that more than eighty percent of the RyR is located on the cytosolic side and is shaped like a rectangular prism (29 × 29 × 13 nm) with several cavities (Radermacher et al., 1994; Wagenknecht and Radermacher, 1995). With the significant sequence similarity between the three isoforms, it is expected that they all have a similar three-dimensional structure and comparable gating movements. All three isoforms exhibit a biphasic response to free Ca2+ and can participate in CICR in permeabilized systems. All isoforms of RyRs show diverse sensitivities to Ca2+-dependent inactivation and activation. Generally, ∼0.3 µM to 10 µM Ca2+ activates RyR, and millimolar concentrations of Ca2+ inhibit RyR activity (Fill et al., 2000). This sensitivity to Ca2+ keeps the RyRs closed at the resting physiological concentration of Ca2+, which is about 100 nM. More details on the structure and function of RyR2 are discussed in Section 1.3. The macromolecular complex of RyR within the SR Ryanodine receptor channels, together with a number of other molecules in the SR, form a macromolecular complex. On the SR membrane, junctin and triadin, whose N-terminus extends a short way into the cytosol and CSQ, the SR-bound Ca2+ storage molecule with a high affinity but   9 low capacity Ca2+ binder, form part of the macromolecular complex (Wang et al., 1998). CSQ interacts with triadin and junctin to affect RyR function and regulate CICR. Triadin (Guo et al., 1996) and junctin (Jones et al., 1995) have comparable structures.  Their C-terminal domains bind to CSQ and RyR2 (Figure 1.4). The cardiac muscle of junctin-null mice seems to have no significant modifications in their cellular structure. These mice die immature, as young as five weeks old, due to an increase in the SR Ca2+ release and to arrhythmias related to delayed-after-depolarizations (Yuan et al., 2007).  Triadin-null mice appear to grow normally, except they have fewer and smaller junctions, and less RyR2 compared to wild type (Chopra et al., 2009). Characterization of this animal model showed a complete absence of junctin, reduced expression of CSQ and lower co-localization of RyR2 and Cav1.2. Furthermore, these mice exhibited a risk of an enhanced SR Ca2+ leak associated with catecholaminergic polymorphic ventricular tachycardia (CPVT).  It has been reported that patients with a mutation in triadin also suffer from CPVT, which can lead to sudden cardiac death (Roux-Buisson et al., 2012).  Calsequestrin is a major Ca2+ buffer and storage protein located in the SR lumen of cardiac (CSQ2) and skeletal (CSQ1) muscles. CSQ is highly acidic and has up to 50 Ca2+ binding sites, each of which is formed by the clustering of three or more acidic residues (Beard et al., 2004). CSQ’s affinity for Ca2+ is between 1 µM to 100 µM, which is species and isoform dependent. At the diastolic luminal Ca2+ concentration (estimated to be 1-1.5 mM (Gyorke et al., 2007)) CSQ forms a polymer that is anchored to RyR either directly or via triadin and junctin (Jones et al., 1998).  Other than being the major Ca2+ buffer and storage protein, CSQ also regulates RyR2 activity. In the presence of junctin and triadin, CSQ inhibits Ca2+ release at the luminal diastolic free Ca2+ concentration. Whether changes in Ca2+ concentration in the SR lumen or   10 conformational changes in CSQ (dissociation from jSR) activate Ca2+ release through RyR2 (increase RyR2 open probability) is not clear, although there is sufficient evidence available to support both mechanisms (Gaburjakova et al., 2012). Mutation of the human CSQ2 gene has been linked to CPVT and sudden cardiac death (Faggioni and Knollmann, 2012). Studies of CSQ2 null mice have shown changes in the myocytes’ cellular structure, in which the SR appears swollen. These mice are also subject to CPVT, most likely due to a reduction in control of the RyR2 open probability that results in an increased SR Ca2+ leak (Knollmann et al., 2006).  1.2.2 L-type calcium channel  Cav1.2 is composed of four different subunits: the α1C subunit forms the Ca2+ selective pore and contains binding sites for the majority of its regulatory proteins, while the extracellular α2δ subunit, the γ subunit and the β subunit are all involved in trafficking, anchoring and in regulatory functions (Catterall, 2011). Immunofluorescence studies in rat cardiac myoctes have indicated that more	  than	  75% of Cav1.2 clusters are located within dyads (Scriven et al., 2010). They are tinier than the RyR2 clusters and appear to be located over the centres (Scriven et al., 2010). Some of the functional analyses suggest a tight physical association between RyR2 and Cav1.2. In these studies, the Ca2+ spark frequency in permeabilized cells are decreased by antagonists and increased by agonists in the absence of a Ca2+ gradient or flux through the channel (Copello et al., 2007; Katoh et al., 2000). This implies that Cav1.2 affects RyR2 activity via a direct physical interaction or via intermediary proteins. In validation of this view, other groups observed direct binding of the C-terminus of the α1C subunit to RyR2 (with a high affinity, 6 nM). When the C-terminus was injected into atrial cells (without Cav1.2 adjacent to RyR2), the Ca2+   11 spark frequency increased, confirming that this peptide directly affects RYR2’s open probability (Mouton et al., 2001; Woo et al., 2003).     1.2.3 The sodium-calcium exchanger NCX is a secondary active transporter, translocating three Na+ ions in exchange for one Ca2+ ion across the plasmalemma, and is the main route for Ca2+ extrusion from all myocardial cells (Hryshko, 2011). The exchanger is reversible, with the direction of transport being determined by the local Na+ and Ca2+ gradients as well as the membrane potential. The reversibility has led to the proposal that under some circumstances NCX can bring Ca2+ into the cell and directly affect, or initiate, ECC. The strongest support for this hypothesis has come from the cardiac-specific NCX knock-out (KO) mouse (Henderson et al., 2004). A comparison of ECC in the KO and wild-type (WT) has demonstrated that current through the voltage-gated Na+ channel (INa) increases the local Na+ concentration reversing NCX and augmenting ECC, an effect completely absent in the KO mice. In these experiments, NCX could not initiate ECC, but the Ca2+ it brought into the cell ‘primed’ the RyR2 and acted synergistically with Ca2+ entering through Cav1.2 (Larbig et al., 2010; Torres et al., 2010). These data indicate that voltage-gated Na+ channels and NCX have functional access to the dyadic cleft, but cannot discern whether they are nearby or physically within it. Immunofluorescence studies in ventricular cells have provided conflicting results, with co-localization values for RyR2-NCX reported as varying from 3-10% (Dan et al., 2007; Scriven et al., 2000b) to ∼42% (Jayasinghe et al., 2009). Triple co-localization experiments have been performed in rat atrial cells, and demonstrated a statistically significant number of Cav1.2-RyR2-NCX triplets including ∼13% of the NCX and RyR2 clusters (Schulson et al., 2011). These experiments also demonstrated a much tighter association between RyR2 and   12 Cav1.2 than between Cav1.2-RyR2 and NCX, suggesting that NCX is at the periphery of the Cav1.2-RyR2 clusters. These co-localization values are also consistent with NCX currents in pig myocytes that have demonstrated that 10–15% of NCX current (INCX) is in or near the dyadic cleft (Acsai et al., 2011). Unpublished results from our laboratory indicate that the molecular architecture of the ventricular dyads is similarly diverse.  1.2.4 Caveolin 3  Caveolae are plasma membrane invaginations that are enriched in cholesterol, sphingolipids, and the marker protein, caveolin (Palade 1953). Three caveolin genes have now been identified (Cav-1, -2, and -3) (Song et al., 1996). Cav-1 and Cav-2 are abundantly co-expressed in a variety of cell types, e.g. adipocytes, fibroblasts, endothelial cells and smooth muscle cells. In contrast, Cav-3 is the only caveolin family member expressed in striated muscle cells (cardiac and skeletal). Cav-3 has been shown to associate with the developing t-tubule system (Parton et al., 1997), and it was originally thought that t-tubules grew through an extension of caveolae. The expression of Cav-3 has been recently reported to be reduced in culture after 72h in rat cardiac myocytes, but not in mouse myocytes. This has been reported to be important since it makes mouse a better model to examine Cav-3 function and distribution in cardiac cells (Pavlovic et al., 2010). Caveolae have also been shown to function as "pre-assembled" signaling complexes through the compartmentalization of signaling molecules, ion channels and transporters that interact with the caveolin proteins. In the ventricular myocyte, a variety of signaling molecules such as β2-adrenergic (β-AR), M2-cholinergic, and A1-adenosine receptors, G-proteins, PKA, PKC and eNOS co-fractionate with caveolae, and their location in caveolae is important for their function (Champion et al., 2004; Takimoto et al., 2005). In addition, all of the   13 sarcolemmal proteins that are important for regulating ECC in cardiomyocytes have also been associated with Cav-3, including Cav1.2, Ca2+-ATPase, Na+/K+-ATPase, Na+ channel, and NCX. These observations are largely indirect, depending on cell fractionation and immunoprecipitation, sometimes using over-expression in heterologous systems, but there are some direct colocalization analyses. These findings, which showed the development and organization of t-tubules and the coordination of second messenger systems with ion channels and transporters, suggest that caveolae are critical determinants of cellular morphology and molecular architecture critical for regulating EC coupling. Previously published data from our lab (Scriven et al., 2005), using indirect immunofluorescence, demonstrated a population of RyR2 clusters without adjacent Cav1.2 but instead colocalized with Cav3. Since I cannot find RyR2 anywhere outside of a structure resembling a couplon, I have hypothesized that these smaller RyR2 clusters are adjacent to Cav-3. If true, there are at least two types of couplons in a ventricular myocyte, a Cav1.2 couplon and a Cav3 couplon. Cav3 is the muscle-specific isoform of caveolin family, localized on sarcolemma, and the major isoform expressed in cardiac cells. The Cav3 couplons are not randomly distributed, instead our data indicate that they are positioned significantly closer to the Cav1.2 couplons than the Cav1.2 couplons are to each other (Scriven et al., 2010).  1.2.5 Junctophilin  An essential component of the dyadic cleft is the molecule junctophilin 2 (JPH2), one of a family of four isoforms which are present in the junctional membrane complexes of other excitable cells such as skeletal muscle (JPH1) or brain (JPH3 and 4) (Nishi et al., 2003;   Takeshima et al., 2000). JPH2 is a 74-kDa molecule, which spans the cleft from the plasmalemma   14 to the SR, and plays the central role in the development and maintenance of that space. The N-terminal end touching the plasma membrane has 8 MORN (Membrane On Receptor Nexus) sequences while the other end, the transmembrane domain, is embedded in the SR. The MORN sequence is highly conserved across species (Garbino et al., 2009) and seems to be responsible for binding both to lipids and proteins in the t-tubular membrane. The section spanning the dyad cleft consists of an alpha helix as well as a divergent region that is associated with the characteristics of each isoform. There are a number of putative phosphorylation sites on JPH2, but none has been associated with a particular function (Garbino et al., 2009). In skeletal muscle, there is evidence that JPH1 is linked to both Cav1.1 and RyR1 (Golini et al., 2011) but the exact relationship in cardiomyocytes is unclear; super-resolution images of JPH2 and RyR2 show differing distributions that vary from dyad to dyad, ranging from each RyR2 having a JPH2 adjacent to it, to having the JPH2 at the periphery of the dyad surrounding all of the RyR2 within it (Jayasinghe et al., 2012). In mice in which the knock down of JPH2 was achieved using tamoxifen-induced siRNA, there was a 40% reduction in the number of dyads (van Oort et al., 2011). The width of the remaining dyadic clefts was, on average, no different than control, but displayed more variance, even within a single dyad (1.96 nm2 in siRNA-treated mice vs. 1.21 nm2 in the controls). The functional consequences were a significant reduction in Cav1.2 co-localization with RyR2 and in the amplitude of the Ca2+ transient, but no change in the Ca2+ current. These results suggest that JPH2 is not only responsible for maintaining the dyadic structure but also keeps the Cav1.2 positioned opposite the RyR2, a role similar to that in skeletal muscle. This would be consistent with recently reviewed functional and structural evidence pointing to a much closer physical association between Cav1.2 and RyR2 than previously thought.    15 1.3 Ryanodine receptor’s structure and function The ryanodine receptor’s name is adapted from a South American plant Ryania speciosa (Rogers et al., 1948).  Ryanodine, the active alkaloid component of this plant, targets RYRs and locks these channels into a subconductant state at low concentration and blocks them completely at 100 µM concentration (Meissner, 1986). RyRs were visualized in electron microscopy studies of muscle ultrastructure. They appear as electron dense lumps, named “feet”, between the SR and sarcolemma in dyads (Franzini-Armstrong, 1970; Lai et al., 1988; Inui et al., 1987). The three isoforms share ∼65-70% sequence identity and the largest difference is in three “divergent regions”, known as DR1 (residues 4210–4562 in RyR2), DR2 (residues 1353–11397), and DR3 (residues 1852-1890). These regions are important since they are thought to be largely responsible for the differing properties of the isoforms. Non-mammalian vertebrates express two isoforms, RyRα and RyRβ, while only one isoform is expressed in invertebrates, including nematodes, flies, and lobster (Lanner et al., 2010). The main opening trigger for RyR is the same ion it permeates, Ca2+. In CICR of cardiac myocytes, Ca2+ sensors within RyR2 bind Ca2+ that is released through Cav1.2, causing channel opening, resulting in the release of Ca2+ from the SR. However, as Ca2+ levels in the dyadic cleft rise, Ca2+ binds to the RyR2 inactivation site, preventing further Ca2+ permeation. This has been interpreted as evidence that there are multiple sites for Ca2+ to bind with different affinities on the cytosolic side of the channel. RyRs also sense the Ca2+ concentrations in the SR lumen. In conditions where the SR is overloaded with Ca2+, RyRs open spontaneously in a process recognized as store overload-induced Ca2+ release (SOICR) (Jiang et al., 2004; Palade et al., 1983). More information regarding the RyRs sensitivity to Ca2+ has been discussed in Section    16 1.3.1 Ultrastructure of the RyR Cryo-EM studies with resolutions around 1nm describe the overall structure of the channel (Samso et al., 2005; Di Maio et al., 2007; Ludtke et al., 2005). RyRs have a mushroom shape with a large cap located in the cytoplasm and a stalk that crosses the SR membrane. The TM region measures 12 × 12 × 6 nm, whereas the myoplasmic area (MY) measures 29 × 29 × 13 nm. To enable a discussion of RyR tetramer structure, different parts have received names, including “clamps,” “handles,” and a “central rim” that surrounds a central cavity. Fifteen globular subregions have been identified on the myoplasmic region. Based on the highest resolution map, subregions 5, 6, 7a, 7b, 8a, 8b, 9 and 10 are located on the clamps at each corner, subregions 3 and 4 are located on the handles and subregion 1, 2a and 2b are located on the central rim. Subregions 11 and 12 are located at the columns that connect the MY region of RyR to TM (Serysheva et al., 2008).  RyRs transport ions through the pore, located at the centre of the central cavity. The clamps located at the corners of the myoplasmic side of the tetramer are regions that are thought to be for interaction with the neighboring channel (Samso et al., 2005; Sharma et al., 1998) (Fig. 1.3).  Most cryo-EM studies have been performed on RyR1 due to its great abundance on the SR and its relatively easy purification. The overall shape is very similar for all RyR isoforms (Sharma et al., 1998). Movements associated with channel gating and coupled gating The recent work of single-particle cryo-EM has revealed the structural transition associated with the RyR1 gating. The three isoforms have near identical three-dimensional structures, as expected from the large sequence similarity, so comparable gating movements are expected from RyR2. Most of the structural differences between the open and closed states, other   17 than the widening of the pore in the open state, are in the clamps and transmembrane regions. Compared to the closed state of RyR1, the clamp regions are about 0.8 nm closer to the surface of the SR and subregion 2 in the central rim moves upward and away from SR membrane in the open state (Samso et al., 2009). Since most of the changes happen far from the pore area, RyRs are allosteric proteins and it is possible to affect their function by interrupting any of these movements.  The two-dimensional crystallization of RyR1 suggests that subunits 6 and 9 in the clamp region (Fig 1.3 C) are responsible for inter-protein interaction (Yin et al., 2005b). As the clamp region undergoes considerable motions during opening and closing (Samso et al., 2005), motions in one tetramer could be spread to neighboring tetramers through physical interactions. These findings support the hypothesis that communication between the tetramers within the array are also regulated by allosteric mechanisms, and that multiple RyR tetramers can open and close in a concerted form. This phenomenon, at the functional level, was introduced as coupled gating in RyR2 (Marx et al., 1998) and RyR1 (Porta et al., 2012b). Functional studies of RyR1 in lipid bilayer suggested that the appearance of coupled RyR1 tetramers is caused by Ca2+ flux from the lumen to cytosol (Laver et al., 2004; Liu et al., 2010). Others however have shown that synchronicity among either RyR1 or RyR2 is not dependent on Ca2+ flux because coupled events were observed when Ca2+ was replaced with Ba2+ as the charge carrier (Gaburjakova and Gaburjakova, 2008; Marx et al., 2001a;). Ba2 + has been used as a replacement for Ca2 + since it is a competitive non-agonist and by itself, does not activate RYR2 channels from the cytosolic site (Gaburjakova and Gaburjakova, 2006). These observations indicated that coupled gating is not dependent on Ca2 + flux through RyR. It is not clear whether the difference in the results of these studies are due to difference in experimental method or whether Ba2 + can compete with Ca2 + on   18 binding sites. In both cases, it is clear that physical communications between tetramers, coupled gating, modifies RyR function and has a role in controlling Ca2 + release during ECC. Other studies of RyR2 in lipid bilayer, comparing coupled and single channel behaviors, also showed that the communication between tetramers significantly affects channel regulation and coupled gating behaviors (Gaburjakova and Gaburjakova, 2014).   Figure 1.3 CryoEM map of RyR1 tetramer as viewed from the myoplasm (A), and from the side (B). TM, transmembrane stalk; MY, myoplasmic rectangular prism, yellow circled regions highlight the domains forming the clamp, dashed yellow highlight the central domain. (C) Two tetramers showing the expected interaction via the clamp domain. (Cryo-Em map of RyR1 channel (EMDB 1606) was uploaded from (Pettersen et al., 2004)   10 9 6 6 10 9 Handle'A' B'C'2 9 10 6 ''''''''''MY'TM''''''''''''  19 1.3.2 RyR arrays The dyadic cleft was originally thought to be filled by tetramers, but new imaging technologies have proven otherwise. Super-resolution immunofluorescence microscopy of rat ventricle myocardium (30 nm resolution) has shown that RyR clusters are much smaller than previously thought. In a dyad, the mean number of RyR2s is around 13, or around 21 in a ‘supercluster’ containing small groupings with edge-to-edge distances of approximately 100 nm (Baddeley et al., 2009). This is vastly different from earlier estimations, based on 2D EM and the assumption of a filled dyadic cleft: 267 RyR2s in a rat ventricular dyad, 90 in dog and 128 in mice (Franzini-Armstrong, 1999). Electron tomographic data from Hayashi (Hayashi et al., 2009) produced similar results and showed that only about 78% of the dyadic cleft was occupied by RyR2 (Hayashi et al., 2009). Both super-resolution immunofluorescence and electron tomography identified isolated RyR2 within the cleft that likely correspond to ‘rogue’ RyRs (Sobie et al., 2006). It has been suggested that these isolated channels may have a different response to local Ca2+ (Sobie et al., 2006).  All together, it is clear that cluster size varies extensively.  Array arrangement seems to be an inherent property of the RyR tetramers since purified RyR1s are capable of self-assembly into 2D arrays either on the surface of lipid bilayers (Yin et al., 2005b) or in buffers (Yin and Lai, 2000). However, in 2D crystallization studies of RyR1, and in a high free Mg2+ concentration, a side-by-side array was formed that precluded allosteric interactions (Yin et al., 2005a). The checkerboard array only formed in nominally-free Mg2+ concentrations. In skeletal muscle triads, RyR1 forms a checkerboard array whereby each tetramer contacts its neighbors through subregions 6 and 10 in the clamp region (Ferguson et al., 1984) (Figure 1.3 C). Electron micrographs of thin sections of cardiac dyads have indicated a regular spacing between the tetramers, comparable with that of skeletal muscle triads (Franzini-  20 Armstrong, 1999). Due to the structural likeness to RyR1, RyR2 has also been assumed to form a checkerboard array.  None of these studies could determine the distribution of individual tetramers within a cluster. However, results from current study (Chapter 4), using electron tomography produced new insights into the organization of RyR2s within the dyadic cleft.  1.3.3 Regulation of RyR2 The activities of all three RyR isoforms are regulated by a significant number of agents, including ATP, Ca2+, Mg2+, junctin, calmodulin, CSQ, FKBP12, Calmodulin (CaM), triadin, sorcin and many others. Most of these modulators bind to the cytosolic domains of the RyRs except junctin, triadin and CSQ, which are on the luminal side (Figure 1.4). In addition, post-translation modifications such as phosphorylation and oxidation also affect RyR gating. RyRs form a macromolecular complex with regulatory proteins, kinases, phosphatases and cytoskeleton proteins (Bers, 2004; Marx et al., 2000). One of the prerequisites for understanding the mechanisms of action of modulators and modifications is the identity of the sites of interaction in the primary sequence and the location of these sites in the 3D structure. Single-particle cryo-EM has been used to analyze macromolecular interactions of RyR with the modulatory proteins (Serysheva et al., 2007).   To evaluate the functional effect of these regulators a variety of model systems have also been developed over the years. Native and recombinant expression systems and, mostly, in vivo systems have been used to examine the role of these regulators on RyR2 activity.    21 Calcium, magnesium and ATP ATP, Ca2+ and Mg2+ are key regulators of RyRs (Meissner, 1994); Coronado et al., 1994). 1 µM Ca2+ and mM ATP activate RyRs and mM level of Mg2+ has inhibitory effects on the channel (Laver et al., 1997) and (Meissner, 1994). These regulatory effects are different among isoforms, probably due to variable characteristics of ECC in cardiac and skeletal muscles.    RyR2s are initially activated by Ca2+ on their cytosolic side. Then, CICR provides a positive feedback to opening more RyR2s resulting in up to 95% of the Ca2+ store being released (Bers, 2002). Increased Ca2+ level in the dyadic cleft and decreased Ca2+ in the SR lumen are thought to be the principal mechanisms that inactivate and close RyR2s, terminating SR Ca2+ release. SERCA2a pumps the Ca2+ back into the lumen and NCX extrudes Ca2+ from the cell. So during ECC the major cytosolic Ca2+ changes, from 0.1 µM (diastole) to 1 µM (systole), and luminal Ca2+ changes, from 1 mM (diastole) to 0.3 mM (systole), are from entry through Cav1.2 and release through RyR2, and removal by NCX and uptake by SERCA2a. Ca2+ not only has a direct affect on RyRs but also regulates the channels through other regulators, such as CaM and CaMKII.  Early studies measuring RyR2s’ open probability (Po) indicated that cytosolic Ca2+, all by itself, in the absence of other regulators, can increase Po from almost 0 to 0.6 (Xu et al., 1996).  It has been well known that there is at least one high affinity (1 µM) Ca2+ activation site (Zahradnik et al., 2005) and at least one low affinity, Ca2+ (1 mM) inhibitory site (Balog et al., 2001) on the cytosolic side of each RyR2 monomer. Later studies of single channel recordings identified three regulatory sites on the cytosolic side and one on the luminal side of RyR2. However, the exact locations of these sites have not yet been determined (Laver, 2007). It has been suggested that these sites’ sensitivities to Ca2+ are all dependent on their distance from the channel pore; the   22 further it is located, the lower its sensitivity (Laver, 2010). The A-site, which is the closest cytosolic activation site to the pore, has a dissociation constant of 0.9 µM, and I2-site, the cytosolic inhibition site, is located most likely on the clamp region and has a 1.2 µM affinity to Ca2+. The I1-site, which is located furthest from the pore, has a very low affinity to Ca2+ (10 mM).  The only luminal activation site (L-site) in RyR2 was first reported in the early 1990’s (Sitsapesan and Williams, 1994). The luminal Ca2+ site has been characterized and its affinity to Ca2+ was reported to be around 40 µM, which is much lower than the A-site. Even with a very low affinity, luminal Ca2+ can still activate adjacent RyR2 in the absence of cytosolic Ca2+. This is because Ca2+ flow through the pore can also activate RyR2 channels (Laver, 2010). Recently, the exact luminal Ca2+ site (E4872A) of RyR2 has been reported (Chen et al., 2014). It has been shown that the point mutation of this Ca2+ site entirely pulls down luminal, but not cytosolic, activation of RyR2 (Chen et al., 2014).  All together, cytosolic and luminal Ca2+contribute to RyR2’s activation and their absence nearly inactivates RyR2 channels (Laver, 2010).  RyR closure is hypothesized to be largely the result of Ca2+ depletion in the local jSR. That idea arose from lipid bilayer experiments demonstrating that the myoplasmic sensitivity of RyR to Ca2+ was a function of luminal Ca2+ (Gyorke and Gyorke, 1998). This has been supported by single cell studies using low affinity Ca2+ indicators to directly monitor SR Ca2+, which clearly demonstrate a reduced, but highly localized, drop in SR Ca2+ (a Ca2+ blink), accompanying the Ca2+ spark (Brochet et al., 2005). More recently, dual recordings of myoplasmic and SR Ca2+, performed simultaneously, confirm that Ca2+ sparks and Ca2+ blinks closely mirror one another in both space and time (Brochet et al. 2011). There is therefore considerable evidence to support luminal Ca2+ depletion in the local jSR as one factor controlling RyR excitability. However, the   23 SR lumen can be refilled ~5 times faster than the spark amplitude recovers, so the sensitivity of the RyR recovers more slowly than the SR Ca2+content (Brochet et al., 2005).  Just like Ca2+, cytosolic and luminal Mg2+ have a role in regulating RyR2 activity. The cytosolic inhibitory effect of Mg2+ was recognized shortly after RyR channels were identified, and its luminal inhibitory effect was just recently characterized (Laver and Honen, 2008). Mg2+ inhibits RyR2s by competing with Ca2+ binding sites (Smith et al., 1986), but is an antagonist that can directly close RyR even in the presence of other activators (Laver et al., 2004). Recently, the Mg2+ affinities for the Ca2+ binding sites, both in the cytosol and luminal sites, have been measured; Mg2+ binds to the A-site with an affinity of approximately 60 µM and to the L-site at ~40 µM. At the luminal side, Mg2+ inhibits RyR by occupying the Ca2+ activation site (L-site) and at the cytosolic side, Mg2+ binding to the A-site can close the channel even when the luminal site is occupied by Ca2+ (Laver, 2010). The physiological concentration of Mg2+ is around 1 mM in both cytosol and lumen. RyRs are permeable to both Mg2+ and Ca2+, so Mg2+ flow can also inhibit RyR activity. Overall, channel gating is triggered by a mutual action of all cytosolic and luminal sites. ATP at millimolar level can activate RyR2s but only in the presence of Ca2+. It has been suggested that ATP stabilizes RyR2s in open conformation associated with high affinity binding sites (Laver, 2007). FKBP 12 FKBP12 and FKBP12.6 (also known as calstabin 1 and 2, respectively) belong to the family of immunophilin proteins that physically interact with RyRs. Three-dimensional reconstruction of RyR locates the FKBPs binding site on subdomains 9, 5 and 3 (Samso et al.,   24 2006). FKBP12 binds to RyR1 and FKBP12.6 binds to RyR2 in the same orientation with a stoichiometry of 4 FKBP per tetramer (Cornea et al., 2010). FKBP12.6 in cardiac and FKBP12 in skeletal muscles are thought to stabilize the closed state of the RyR channels (Marx et al., 2000). Also it has been suggested that FKBP12 has a key role in coupled gating between the channels (Ondrias et al., 1998). Abnormal gating behaviour has been reported in FKBP12 KO mice (Shou et al., 1998) and removal of FKBP12 by immunosuppressive drugs (Rapamycine or FK506) (Marx et al., 1998). However, single channel recordings of RyR2 revealed that the removal of FKBP12.6 has no effect on the channel functions (Barg et al., 1997). It has been suggested that PKA phosphorylation of RyR2 causes the dissociation of FKBP12.6, which leads to enhanced channel opening and abnormal gating (Marx et al., 2000). However, this hypothesis has been challenged by other groups showing that phosphorylation by PKA has no effect on binding of either FKBP12 or 12.6 to RyR2 (Guo et al., 2010). A recent study, using Western blot and [3H] ryanodine binding assays, also showed a significant difference between species in the molar ratio of RyR2-FKBP12.6 and RyR2-FKBP12. In normal heart in mouse, pig and rat, only 35-45% of FKBP12.6 sites whereas in rabbit, 0% are saturated. This indicates that saturation of RyR2 by FKBP12.6 is not critical for normal RyR2 function (Zissimopoulos et al., 2012). It is possible that the difference in species examined and preparations used underlie the inconsistencies between the above studies. Calmodulin  CaM, one of the key regulators of RyRs, can associate with the tetramer in both Ca2+ loaded and apo- conditions and fine-tune the effect of Ca2+. The functional effect of CaM depends on both the RyR isoform and the Ca2+ concentration. At low Ca2+ levels, it activates RyR1 but   25 inhibits RyR2. However at high Ca2+ levels, CaM can inhibit both RyR1 and RyR2 (Ikemoto et al., 1995). In rat cardiac myocytes the cellular concentration of CaM is around 2 to 6 µM, depending on the species (Maier et al., 2006). In normal physiological conditions, CaM binds to more than 70% of RyR2 with a high affinity and inhibits SR Ca2+ release. This is important because in heart failure there is a decrease in CaM and RyR2 association, which may be one of the factors involved in the SR Ca2+ leak (Yang et al., 2014). Phosphorylation RyRs are the target for several kinases (Ca2+-CaM-dependent protein kinase II (CaMKII), PKG, PKA) and phosphatases (PP1, PP2A). Some of these kinases are anchored to RyRs by scaffolding proteins, allowing for specific regulation.  The classical physiological stress pathway, “fight or flight “ response, activates the sympathetic nervous system that results in faster and larger Ca2+ transients (Bers, 2002). Epinephrine and norepinephrine bind to β2 adrenergic receptors (β-AR) on the plasma membrane of cardiac myocytes and stimulate adenylyl cyclase (AC), which generates cyclic AMP (cAMP). cAMP activates PKA and increases the rate of rise and the magnitude of the Ca2+ transient by increasing the activity of RyR2, Cav1.2, and SERCA via phospholamban. CaMKII, on the other hand, is regulated largely by the intracellular Ca2+ concentration and affects the Ca2+ transient similarly to PKA. CaMKII and PKA regulation of RyR2 function have significant roles in controlling cardiac rhythm and contractility. It has been suggested that there is one functional PKA phosphorylation site, S2808 (S2809 in some species), and a separate CaMKII phosphorylation site, S2814 (S2815 in some species), on each monomer of RyR2’s tetramer (Wehrens et al., 2004), but this is far from clear.    26 Single channel studies on lipid bilayers have been shown that the addition of purified PKA increases RyR2 open probability (Marx et al., 2000). Mutation of RyR2-S2808A, showed an increase in the open probability in some studies (Wehrens et al., 2006) but no effect in other studies (Stange et al., 2003). Also, studies on ventricular myocytes from S2808A mice showed changes in spark frequency in some studies (Shan et al., 2010) but not at all in others (Benkusky et al., 2007). Isoproterenol-induced studies on S2808A mice in vivo and in vitro have shown unaltered (MacDonnell et al., 2008) and altered (Shan et al., 2010) chronotropic and inotropic responses. Hyperphosphorylation, saturation of all phosphorylation sites, of S2808 by PKA has been reported to dissociate FKBP12 in heart failure in some studies (Marx et al., 2000; Wehrens et al., 2006), although this has not been demonstrated in other studies (Guo et al., 2010; Zhang et al., 2012). Some investigations even reported increased activity of RyR2s because of dephosphrylation of S2808 (Liu et al., 2014; Terentyev et al., 2003; Valdivia et al., 1995). Multiple factors can produce these differences, including differences in experimental procedures, reagents, and animal models. Another PKA phosphorylation site, S2030 (S2031 in some species), was identified as the third functional site on RyR2 a few years ago (Xiao et al., 2006). Phosphorylation of this site has been reported to increase RyR2 sensitivity to luminal Ca2+ in some studies (Xiao et al., 2006), however other studies failed to validate a functional effect on RyR2 (Huke and Bers, 2008; Wehrens et al., 2006).   Phosphorylation by CaMKII increases RyR2 open probability in single channel recording in bilayers (Wehrens et al., 2004) and increased spark frequency in ventricular myocytes (Respress et al., 2012).  Studies of CaMKII knockout and transgenic animal models have shown changes in spark frequency that are due to altered phosphorylation at S2814 in RyR2 (Ling et al., 2009; Maier et al., 2003). Also, S2814A mutation inhibits most of the CaMKII effects on RyR2,   27 implying that S2814 could be the major, but not the only, CaMKII phosphorylation site (Wehrens et al., 2004). Another interesting observation was that in S2808A knock in mice, S2814 was reported to be hyperphosphorylated, which implied a cross-talk between phosphorylation sites (Benkusky et al., 2007). In vitro studies on phosphorylation domains of RyR2 have revealed several phosphorylation sites other than S2808, S2814 and S2830. Overall five phosphorylation sites have been reported for PKA: S2814, T2810, S2811, S2797 and S2808, and four phosphorylation sites for CaMKII:  S2814, S2808, S2811 and T2876 (Yuchi et al., 2012). It is important to note that based on this in vitro study both PKA and CaMKII can phosphorylate S2808, S2811 and S2814. Phospho-proteome investigation of different mice organs demonstrated that all these sites, except T2876, could be phosphorylated by PKA and CaMKII in vivo (Huttlin et al., 2010). However, in the physiological state within cardiomyocytes, it is not clear, which kinase phosphorylates which sites. RyR2 phosphorylation and cardiac disease Phosphorylation of RyR2 channels are important in pathophysiological conditions such as heart failure (Marx et al., 2000; Wehrens et al., 2006), as well as in atrial and ventricular arrhythmogenesis (van Oort et al., 2010). Hyperphosphorylation of RyR2 increases channel open probability, resulting in diastolic Ca2+ leak. This has two distinct and undesirable effects: 1) A decrease in the luminal Ca2+ level producing a smaller Ca2+ transient and impaired contraction that can result in cardiac failure and 2) Diastolic Ca2+ leak can trigger DADs (delayed afterdepolarization) and arrhythmias.      28 There have been several studies indicating that in heart failure, RyR2 are hyperphosphorylated by PKA (Marx et al., 2000). CaMKII phosphorylation also has been reported to contribute to cardiac arrhythmogenesis and heart failure (Fischer et al., 2013; Guo et al., 2006; Respress et al., 2012), but the relative importance of Ca2+ leak produced by PKA versus CaMKII in heart failure is unknown. However there are several laboratories trying to differentiate the role of these two kinases by developing various animal models, and by examining cardiac remodeling and patients with heart failure (HF).  One of the first examinations of altered RyR2 phosphorylation in patients with HF postulated that the increase in S2808 phosphorylation by PKA reduced FKBP12.6 binding to RyR2 resulting in diastolic Ca2+ leak (Marx et al., 2000) (Wehrens et al., 2006).  They also demonstrated that the S2808A knockin mice show limited cardiac dysfunction in post-myocardial ischemia (MI) hearts (Marx et al., 2000). Most, but not all, of the other studies confirmed a highly phosphorylated S2808 in HF in different species (Marks, 2013).  Another group using the same model (S2808A) argued that S2808 phosphorylation site is not functionally relevant for MI cardiac function and “fight or flight” response (Zhang et al., 2012).  There have been other reports indicating that S2808 is already phosphorylated (60-80%) in the basal physiological state (Ai et al., 2005; Benkusky et al., 2007; Carter et al., 2006; Fischer et al., 2013; Xiao et al., 2006) and increased slightly in HF (Ai et al., 2005) or failed to detect any rise in phosphorylation level in HF at this site (Xiao et al., 2005). S2030, another PKA phosphorylation site, has also been identified to be more phosphorylated in HF (Xiao et al., 2005). Overall phosphorylation of RyR2 increases in HF and it is not clear which kinase is more important, PKA or CaMKII, although some groups conclude that S2814 phosphorylation by CaMKII is more important in HF (Bers, 2012).    29 Phosphorylation of RyR2 is also reported in CPVT, a genetic condition distinguished by stress or exercise induced arrhythmias and sudden cardiac death (Priori et al., 2001). Over half of the CPVT cases are the result of RyR2 autosomal-dominant missense mutations, and more than 150 mutations has been reported (Priori and Chen, 2011). At the cellular level, enhanced β-AR activity due to stress or exercise triggers CaMKII and PKA, resulting in RyR2 phosphorylation, SR Ca2+ leak and arrhythmias in CPVT.  The majority of the studies on CPVT mutations reported unaltered baseline phosphorylation levels on S2808 and S2814 (Shan et al., 2012) and no CPVT mutation has been found near these two phosphorylation sites in RyR2 (Yuchi et al., 2012) but 11 have been reported in RyR1 (Yuchi et al., 2012).  Mechanisms underlying different effects of PKA and CaMKII on RyR2 gating  RyR2 can be phosphorylated directly by CaMKII, which causes both diastolic SR Ca2+ leak and sensitizes the RyR2 to Ca2+-induced Ca2+ release during ECC. It has been suggested that RyR2 phosphorylation by CaMKII increases spark frequency without requiring a change in the SR Ca2+ content. There is general agreement on the effects of CaMKII on RyR2. However the effect of PKA phosphorylation on channel gating is controversial. The mechanism suggested by one group is that PKA phosphorylation of RyR2 at 2808 plays an important functional role for synchronizing Ca2+ release and sensitizing RyR2 to the luminal Ca2+ level. This suggests that there is a direct link between RyR2 phosphorylation and SR luminal Ca2+ sensing by PKA (Ullrich et al., 2012). The other mechanism for which there is empirical evidence is that RyR2 phosphorylation by PKA might speed up the Ca2+ release process, but does not increase the Ca2+spark frequency or diastolic SR Ca2+ release without an increase in the SR Ca2+ content (Bers, 2012).   30   Figure 1.4 Modulators of RyR2.  A list of inhibitors and activators of RyR2, as well as binding partners in the myoplasmic (MY) area and luminal (TM) areas. (Cryo-EM map of the RyR1 side view adapted from       Ca2+%Triadin%%%%%%Junc.n%CSQ%CSQ%Low%concentra.on%of%Ryanodine%Low%Ca2+%Concentra.on%Caffeine%PKA%CamKII%ATP%%%%%%High%Ca2+concentra.on%%%High%Ca2+concentra.on%CaM%FKBP12/12.6%High%concentra.on%of%Ryanodine%Tetracain%Mg2+%%%%%%%%%Low%Ca2+concentra.on%%%%%Increase(Ca2+(release( Inhibit(Ca2+(release(MY%TM%  31 1.4 Research hypotheses and aims 1.4.1 Chapter 2 Hypothesis: RyR2 is in cSR and is adjacent to caveolae. Aim: Using transmission electron microscopy and tomography examine ventricular muscle and quantify the distribution of RyR2 in cSR vs dyads. Rationale: Diffraction-limited immunofluorescence had identified a population of RyR2 that colocalized with caveolin-3, but not with Cav1.2. 1.4.2 Chapter 3 Hypothesis: RyR2s are not distributed in a regular array. Aim: To determine whether the distribution of RyR2 in a dyad examined by tomography could be fit with a regular array of tetramers. Rationale: Preliminary data suggested that the tetramers were not distributed in a regular array. 1.4.3 Chapter 4 Hypothesis: RyR2s are not distributed in a regular array and their relative positions are not fixed. Aim 1: To determine whether the distribution of RyR2 in a dyad examined by tomography could be fit with a regular array of tetramers. Aim 2: To quantify the distribution of the tetramers in rat and human ventricular myocytes. Aim 3: To determine the effects of physiologically and pathologically relevant factors on the distribution. Rationale: Our analysis of an IMOD image suggested that the tetramers were not in a regular array. In vitro analysis of RyR1 tetramers demonstrated that the RyR2 distribution might not be fixed.    32 2. Axial tubules of rat ventricular myocytes form multiple junctions with the SR 2.1 Synopsis Ryanodine receptors are largely on jSR adjacent to the transverse tubules and on the cell surface near the Z-lines, but some RyR are on jSR that is adjacent to axial tubules. Neither the size of the axial junctions nor the numbers of RyR they contain has been determined. RyR may also be on corbular SR and on free or network SR. Since determining and quantifying the distribution of RyR is critical for both understanding and modeling Ca2+ dynamics, I have investigated the distribution of RYR in healthy adult rat ventricular myocytes using electron microscopy, electron tomography and immunofluorescence. We only found RyR in three regions, in couplons on the surface and on transverse tubules, both of which are near the Z-line, and in junctions on most of the axial tubules; axial junctions. The axial junctions averaged 510 nm in length, but occasionally spanned an entire sarcomere. Numerical analysis showed that they contain as much as 19% of the cell’s RyR. Tomography confirmed the axial junction’s architecture, which is indistinguishable from junctions on t-tubules or on the surface, and revealed a complexly structured tubule whose lumen was only 26 nm at its narrowest point. RyR on axial junctions co-localize with Cav1.2 suggesting they play a role in excitation-contraction coupling.  2.2 Introduction The sub-cellular distribution of RYR in a myocyte determines the size, shape and duration of the whole cell Ca2+ transient, and creates microdomains where the local Ca2+ concentration is far greater than the cellular average. If we are to model the Ca2+ transient, and to understand local   33 physiological control of Ca2+-dependent processes, we must know where the RyR are located, as well as the organelles and other molecules to which they are adjacent. Our current understanding is that most RYRs are located in clusters that are only found in specialized regions of the SR; the junctional and corbular SR. The junctional SR membrane is positioned 10 – 15 nm away from clusters of voltage-gated Ca2+ channels, Cav1.2, that are located either in the sarcolemma or in the transverse-axial tubule system (Carl et al., 1995; Franzini-Armstrong, 1999; Sun et al., 1995). These closely apposed membrane surfaces and their associated proteins create distinct, unmistakable structures in transmission electron micrographs called junctions or couplons. This architecture is essential for normal Ca2+-induced Ca2+ release and excitation-contraction coupling in the heart (Bers, 2002). Most of the couplons in the TATS are formed with transverse tubules that are either at the level of the Z-lines or very near to it and that belong to the transversely oriented regions of the TATS, most properly called t-tubules. In addition, some of the couplons are with axial tubules, but the proportion of axial junctions, their size and the number of RyR they contain, has never been determined.  Corbular SR (cSR) is a RyR-studded sac, about 100 nm in diameter that contains calsequestrin and stores Ca2+ (Jones et al., 1995; Jorgensen et al., 1988). It extends from the SR network into the myoplasm, close to the Z-line, but does not have an adjacent t-tubule or Cav1.2. This is a prominent feature in cells lacking a TATS, such as avian and atrial cells in mammalian heart, but has also been reported in papillary muscle (Jorgensen et al., 1993). A quantitative examination of the extent of RyR on cSR, and the other organelles to which it is adjacent, has not been reported for ventricular muscle.  In addition to these locations, RyR have also been reported in mitochondria (Beutner et al., 2001) and in the free (network) SR surrounding the myofibrils opposite the A-band (Lukyanenko   34 et al., 2007). The content of RyR in mitochondria is very low, if present, but it has been suggested that RyR in the free SR may be fairly abundant and might explain the 15% – 20% of Ca2+ sparks that arise in the A-band, well away from the Z-line and the junctional RyR (Lukyanenko et al., 2007; Shacklock et al., 1995). Little attention has been paid to RyR on axial junctions, even though axial tubules are a well known feature of adult rat cardiomyocytes (Soeller and Cannell, 1999). A quick examination of rat ventricular myocytes reveals that this may be a relatively large proportion of the total RyR and I hypothesize that most axial tubules form junctions with the adjacent SR. To investigate this question I have used a combination of fluorescence microscopy, transmission electron microscopy, and electron tomography.  My results demonstrate that the majority of axial tubules form junctions with the SR and they share all of the structural features of couplons on the plasmalemma and the transversely oriented t-tubules. Axial junctions routinely extend well into the A-band and are occasionally large enough to span an entire sarcomere. Tomography confirmed their structure and revealed an unexpectedly complex morphology of the transverse tubule. Axial junctions are surprisingly numerous, and I estimate that they contain as much as 19% of a ventricular myocyte’s RyR. Immunofluorescent labeling shows that RyR in axial junctions are adjacent to Cav1.2 and could therefore contribute to excitation-contraction coupling.  2.3 Methods All chemicals were purchased from Sigma-Aldrich (Oakville, ON, Canada) unless otherwise stated. Animal handling was done in accordance with the guidelines of the Canadian Council on Animal Care.   35 2.3.1 EM tissue processing Animals were sacrificed with a peritoneal injection of 2 ml of 1000 units of Hepalean (Organon Canada Ltd, Mississauga, ON) and 2.5 ml of sodium pentobarbital (80 mg/100 g; MTC Pharmaceuticals, Cambridge, ON). The hearts were perfused for 10min with physiological saline solution (PSS) followed by a fixative containing 4% paraformaldehyde, 2.1% glutaraldehyde and 4 mM CaCl2 in a 0.1 M cacodylate buffer (pH 7.4; Canemco & Marivac Inc, Lakefield, PQ), for 10 min. The left ventricle or atria was cut into small blocks and the sample immersed in fixative for about 2 hours. To speed up the process, blocks were cyclically microwaved  (2 min on, 2 min off, 2 min on) in a vacuum using a Pelco 3450 laboratory microwave (Ted Pela Inc., Redding, CA), at power 5 (Galvez et al., 2006). Blocks were rinsed and microwaved twice in 0.1 M cacodylate buffer for 40 seconds at power 1 then post-fixed with 1% OsO4 solution (EMS, Hatfield, PA) in the same buffer at power 1, then cyclically microwaved twice. This step removed the cytosolic matrix from the fractured cells at the cracked surface (Tanaka and Mitsushima, 1984). En bloc staining of samples was done with 2% aqueous uranyl acetate (Ted Pella Inc.), cyclically microwaved twice at power 1, and then rinsed three times with distilled water. They were then dehydrated in increasing concentrations of ethanol (50-100% in steps of 10%; microwave 1 min at each dilution on power 3) and embedded in a mixture of Epon and Spurr’s resin. From each block, 2 µm-thick sections were cut with a glass or diamond knife mounted on a Leica Ultracut T (Leica Microsystems, Richmond Hill, ON) and were then stained with toluidine-blue. Suitable sections, on which there were surviving myocardial tissues in a longitudinal orientation, were selected for EM study. Ultra thin sections (80 nm) were cut consecutively from the same block; these were mounted on 0.25% w/v formvar-coated 200 mesh copper grids (EMS, Hatfield, PA) and double-stained with 2% uranyl acetate for 12 min and Reynold’s lead citrate for   36 about 6 min. Sections were investigated using a transmission electron microscope (Hitachi H7600, Hitachi High Technologies America, Schaumburg, IL).  2.3.2 Preparing sections for single-axis tomography From blocks with regions of well preserved SR and tubular system, serial semi-thick (120-130nm) sections were cut using a Leica Ultracut T with a Diatome Ultra 35º diamond knife (Diatome, Hatfield, PA) and collected on 0.5 % w/v formvar carbon coated slot grids. Post-staining was performed with 2% uranyl acetate for 25 min followed by Sato’s lead citrate for 10-12 min (Hanaichi et al., 1986). After post-staining, grids were coated with 7.5 nm colloidal gold particles (BBInternational, Cardiff, UK) for 10 min (Penczek et al., 1992), and then Formvar–coated to enhance sample stability in a high-voltage electron beam.  2.3.3  Image acquisition and tomography Grids were placed in a rotating, high-tilt stage and observed in the Tecnai G2 Sphera TEM (FEI, Eindhoven, Holland) microscope operating at 200kV. A suitable longitudinal junction was imaged at 19000x with serial tilt views from +65° to -65° at 1° intervals using a camera setting of 1024 x 1024 pixels, resulting in a pixel size of 0.90 nm. The tomograms were generated using TIA software (FEI, Eindhoven, Holland). All tilted images were aligned to a common tilt axis using cross-correlation and the volume was reconstructed by a real-space back-weighted projection. Tomograms were displayed and analysed with 3dmod, the graphics component of the IMOD software package (Kremer et al., 1996). The transverse and axial tubules, SR, RyR and CSQ were modeled manually using procedures detailed in Donohoe et al (Donohoe et al., 2006).   37 2.3.4 Fluorescence microscopy Ventricular myocytes were isolated from the hearts of adult male Wistar rats weighing between 200 and 250 grams. Techniques for acquiring myocytes, and for fixation, permeabilization, immuno-labeling, as well as processing, deconvolving and analyzing images from a wide-field microscope have been published (Scriven et al., 2000a; Scriven et al., 2005). An anti-RyR2 monoclonal antibody (Affinity Bioreagents, Golden, CO) and an affinity purified rabbit polyclonal antibody against the pore-forming subunit of the voltage-gated Ca2+ channel, Cav1.2 (Hell et al., 1993) were used. The secondary antibodies were affinity purified and highly cross adsorbed to minimize cross reaction, and were goat anti-mouse conjugated to Alexa 594 and goat anti-rabbit conjugated to Alexa 488 (Invitrogen, Burlington, ON). Images were acquired using an inverted Zeiss Axio Observer microscope equipped with a Plan Apo 63/1.4 objective and EXFO Xcite illumination (Mississauga, ON). All filters are from Semrock (Rochester, NY); exciter FF01-494/20-25, dichroic FF506-Di01-25x36, emitter FF01-536/40-25 for Alexa 488 and exciter FF01-575/25-25, dichroic FF593-Di02-25/36, emitter FF01-624/40-25 for Alexa 594.    2.4 Results The general features common to a junction are shown in the image in Figure 2.1A. The variable features are the size of the junction, which can only be determined from serial sections or tomograms, and the architecture; the SR can form a junction with a portion of the t-tubule, as depicted, or it can encircle all (not shown) or most of the t-tubule (as shown in Figure 2.2E). The architecture changes when the junction is at the cell surface, adopting one of the two configurations shown in the micrographs in Figures 2.1B and 2.1C. Figure 2.1B shows a junction straddling a Z-line. The thin SR connects the two halves of the junction, but the electron dense   38 material in the lumen is confined to the region under the RyR. Seen with equal frequency is a surface junction that is confined to only one side of the Z-line, shown in Figure 2.1C. The centre-to-centre distance between the feet in both internal and surface junctions was the same; 44.3 ± 2.6 nm (mean ± SE; N = 217).    39 Figure 2.1 Transmission electron micrographs of junctions in rat ventricle. A) A dyad on the transverse tubule. Ryanodine receptors (single arrow), sarcoplasmic reticulum (double arrow), mitochondrion (m), Z-line (z). Scale bar = 100 nm. B) and C) Surface junctions. Insets of the indicated regions are magnified 2.5 X. Endothelial cell (ec), interstitial space (is). Scale bars = 500 nm.  We also observed junctions in the interior of the myocyte that are oriented parallel to the cell’s longitudinal axis and perpendicular to the Z-line (Figure 2.2). Axial junctions have all the morphological features as those depicted in Figure 2.1, except they are formed with the axial tubules and they penetrate well into the A-band. The most commonly observed are shown in Figures 2.2A and 2.2B. The former shows an axial tubule that almost spans the sarcomere with a junction that covers only a fraction of its length, while the latter shows a shorter tubule with nearly its entire length covered by a junction. Less common are the axial junctions depicted in Figures 2.2C and 2.2D in which junctions extend almost the entire length of the sarcomere. Axial junctions sometimes originate from a junction on the Z-line, and can appear as a triad; both of these features are seen in the micrograph in Figure 2.2D. Figures 2.2E and 2.2F show axial tubules that do not contain junctions. One of these tubules originates from a junction (2.2E) while the other does not (2.2F). In 45 micrographs encompassing 804 µm2, I observed 187 junctions; 148 were junctions with the t-tubule (79%), 39 were with axial tubules (21%). The majority of the axial tubules, 77%, formed junctions with the SR. The centre-to-centre distance between RYRs on axial junctions was 40.0 ± 0.88 nm, (mean ± SE; N = 94), comparable to that seen in the junctions on the t-tubules and on the surface. The data was obtained from six rats and no differences were observed between the age (200-250 grams) and sex (male) matched animals.   40  Figure 2.2 Longitudinal sections of axial tubules and their junctions. Insets of the indicated regions are magnified 2.5 X. A) and B) Axial junctions that are a fraction of a sarcomere long. C) and D) Axial junctions that are a sarcomere in length E) and F) Axial tubules without junctions.    41 It is clear from Figure 2.2 that the axial tubules and the axial junctions have variable lengths, and that the axial junctions do not always occupy the entire length of the axial tubule. To analyze and quantify these relationships I measured the lengths of the axial tubules and the axial junctions relative to each other, and relative to the sarcomere length. The results are presented in the frequency histograms shown in Figure 2.3, all of which are single distributions (Hartigan and Hartigan, 1985) (p > 0.1) that are non-Gaussian (p < 0.01). I first measured the length of the axial tubules relative to the sarcomere length, Figure 2.3A; sarcomere length was 2.04 ± 0.02 µm (mean ± SE; N=52). The frequency histogram presented in Figure 2.3B demonstrates that the axial junctions occupy a significant fraction of the axial tubules. Finally, the frequency histogram in Figure 2.3C shows that the axial junctions are about one quarter of a sarcomere long, although some are much longer. In cross-section, the axial junctions show the same variable architectures as those with t-tubules, Figure 2.4. The SR can partially or completely encircle the axial tubule (Figure 2.4A), or it can form a junction with one of the flattened surfaces of the junctional SR (Figure 2.4B). In 19 cross-sections I observed a mean of 8.5 RyR per longitudinal junction; range 2-24, median = 7. Atrial cells display surface junctions that are indistinguishable from those in the ventricle, Figure 2.5A, and junctions with transverse axial tubules in those cells that show a rudimentary TATS; an example of an axial junction is shown in Figure 2.5B. All atrial cells, regardless of whether they possess a rudimentary TATS or not, display RyR studded cSR that is readily apparent on or near the Z-lines; an example is shown in Figure 2.5C. The lumen of the SR has electron dense material and the magnified inset shows individual RyR protruding from the membrane into the myoplasm. In contrast, over 200 micrographs of the ventricle failed to find a   42 comparable structure. Instead I observed sacs of SR protruding into the myoplasm that were clearly devoid of RyR, Figure 2.5D.  To more closely view the association between the axial and transverse tubules, I examined a semi-thick section of 120 nm using electron tomography (Figure 2.6). One of the individual slices of the tomogram is displayed in Figure 2.6A, in which the membranes of the transverse and axial tubules, the SR, the electron dense material within the SR and the RyR are clearly visible and have been outlined in different colors, Figure 2.6B. The outlines from all 96 planes form a complete 3D model of the data, Figure 2.6C, which can be seen in a movie sequence in the supplementary material ( A single row of the RYRs was clearly visible, and these, along with the SR, are displayed in Figure 2.6D.   43  Figure 2.3 Frequency histograms. A) The length of each axial tubule was expressed as a fraction of the length of the sarcomere and counted in bins that are one twentieth of a sarcomere long.  N= 52. Mean = 0.56, median = 0.44, mode = 0.26, stnd. dev. = 0.31. B) The length of each axial junction was expressed as a fraction of the length of its axial tubule and counted in bins that are one twentieth of a tubule long. N = 40. Mean = 0.66, median = 0.73, mode = 1.00, stnd. dev. = 0.30. C) The length of each axial junction was expressed as a fraction of the length of the sarcomere and counted in bins that are one twentieth of a sarcomere long. N = 40. Mean = 0.30, median = 0.24, mode = 0.19, stnd. dev. = 0.18   44  Figure 2.4 Cross sections of axial tubules and their junctions. A) The axial tubule is nearly encircled by the junction. The inset is a 2.5 X magnification of the indicated region; the scale bar is 500 nm. B) One surface of the axial tubule forms a junction with the adjacent SR (arrow). Scale bar = 100 nm.      45  Figure 2.5 Atrial RyR and corbular SR. A) Rat atrium. Arrows point to sarcolemma (SL) of adjacent cells; z – z line; m – mitochondrion; Inset is a 3 X magnification of the highlighted surface junction (SJ). Scale bar is 500 nm B) Rat atrium inset is a 3 X magnification of the highlighted axial junction. Scalebar is 100 nm. C) Rat atrium. Arrows point to examples of corbular SR. The inset is a 2.5 X magnification of the indicated region. D) Rat ventricle. The sacs of SR are devoid of RyR. The inset is a 2 X magnification of the indicated region. Scale bars are 500 nm.    46   Figure 2.6 Tomography of a single longitudinal junction. Scale bar = 100 nm. A) One plane of the reconstructed tomogram. B) Tracings of the relevant structures from A. SR (green), individual RyR (red), CSQ (yellow), transverse and axial tubules (blue). C) The drawings from each of the planes forms the 3D model. D) The SR and RyR have been isolated and rotated 70 degrees about the Y-axis.    47 The axial tubule and junctional SR have a relatively simple morphology, unlike the transverse tubule where multiple and interconnected branches are difficult to visualize. To demonstrate the structure of the tubules, I have shown two planes from the tomogram, 72.8 nm apart, in Figures 2.7A and C. Individual sections of the transverse and axial tubules are outlined in Figures 2.7B and D, and for clarity and ease of reference, the different parts of the tubules are numbered 1 through 2.8. The same numbering system was applied to the 3D model, Figure 2.7E, which was rotated about the Y and/or Z axes to produce the images in Figures 2.7F and 2.7G. Over the course of 8.4 nm, the axial tubule widened dramatically from about 20 nm (labeled #1) to about 180 nm (labeled #8). As demonstrated in Figure 2.6C, the change in morphology of the axial tubule was unrelated to the presence of the junction. Tubule 2 is connected to the axial tubule at the point indicated by the arrow in Fig 7E. Tubule 3 is connected to tubule 4 and to the axial tubule as shown in Figures. 7B, E and G. Transverse tubules numbered 4, 5, 6 and 7 connect at the point indicated by the asterisk in Figures 2.7F and G; at this point the tubule is an oval 14 nm wide (Figure 2.7F) and 115 nm long (Figure 2.7G). These four tubules converge and twist, forming an‘X’ whose bottom half is rotated approximately 90 degrees. This can also be seen in Figures. 7B and 7D where a line drawn between the center of tubules 4 and 5 in Figure 2.7B would be roughly perpendicular to a line drawn between the centers of tubules 6 and 7 in Figure 2.7D. Tubule 6 has the smallest diameter and is only 26 nm wide at its narrowest point.   48  Figure 2.7 Tomography of a single longitudinal junction and the transverse tubule to which it is connected. A) and C) Planes 14 and 66 of the tomogram respectively. B) and D) A and C with the tubules numbered and highlighted in blue. Tubules 3, 4, 5, 6 and 7 are transverse tubules, tubules 1and 8 are axial tubules. 3D rendering of the numbered tubules. E) Orientation is the same as in A-D. Arrow points to connection between tubules 2 and 8. F) E rotated 180 degrees about the Y-axis. The asterisk marks the connection between tubules 6 and 7. G) E rotated -90 degrees about the Y-axis, then -45 degrees about the X-axis. The asterisk marks the connection between tubules 4, 5 and 7. Tubule 6 is behind tubule 7 and not visible in this orientation.     49 Lastly, we used dual-label immunofluorescence to determine if axial RyR are adjacent to Cav1.2 (Scriven et al., 2000a; Scriven et al., 2005). A single plane extracted from the 3D image of a cell labeled with antibodies specific for RyR (red) and for the pore-forming α subunit of Cav1.2 (green) is displayed in Figure 2.8Ai (colocalized voxels are white). As expected, both proteins are distributed along the Z-lines, a segment of which is highlighted and magnified in Figure 2.8Aii. Some of the fluorescence is oriented perpendicular to the Z-lines, on axial junctions. We isolated a total of seventy axial and seventy transverse segments from the images; ten of each from seven cells isolated from four rats. Two of the axial junctions isolated from the cell in this image have been highlighted, and single image planes are displayed in the insets. Figure 2.8A iii shows a junction extending almost the entire distance between adjacent Z-lines, and Figure 2.8A iv shows a smaller junction that extends only part way into the sarcomere.  The length of the axial junctions, expressed as a fraction of the sarcomere length, was described by a single distribution (Hartigan’s dip test, p > 0.1), that was non-Gaussian (p < 0.001), Figure 2.8B. Although comparable to that measured using TEM, the mean length measured by fluorescence is significantly larger (Mann-Whitney U test, p < 0.004), which is not surprising given the different resolving powers of the two techniques. A notable difference between the transverse and axial junctions displayed in Figure 2.8A is the RyR on the former that have no adjacent Cav1.2; these are the extra-dyadic RyR (arrow in Figure 2.8Aii). This visual difference is also apparent in the numerical analyses, Fig 8C. The results show that Cav1.2 is colocalized with RyR (black bars) to a greater extent than is RyR with Cav1.2 (grey bars; p < 0.001) on both axial and transverse junctions, which is in agreement with previous observations (Mohler et al., 2005; Scriven et al., 2000a; Song et al., 2006). But, there is a   50 significantly greater amount of colocalization of RyR with Cav1.2 in the axial segments relative to the transverse segments (p < 0.001) due to the lack of extra-dyadic RyR.      51 Figure 2.8 Short and long stretches of axial RyR and Cav1.2. A) Rat ventricular myocyte labeled for RyR (red), Cav1.2 (green). Colocalized voxels are white. i) A single plane of the data set (278 pixels X 406 pixels X 61 planes), scale bar = 5 mm. ii) 4X magnification of the indicated transverse tubule. Image dimensions are 45 pixels X 18 pixels X 4 planes. iii) and iv) 10X magnifications of the indicated axial tubules. Image dimensions are 9 pixels X 18 pixels X 3 planes and 9 pixels X 12 pixels X 3 planes respectively. B) Comparison of the length of the axial junctions measured by fluorescence microscopy (filled bars, N = 70) and electron microscopy (open bars, N = 40) normalized to the count in the largest bin. For the fluorescence microscopy: mean = 0.39, median = 0.32, mode = 1.0, stnd. dev. = 0.23. C) The mean colocalization ± stnd. err. of RyR with Cav1.2 (filled bars) and Cav1.2 with RyR (grey bars). Ten axial tubules and ten transverse tubules were analyzed from each of seven cells. Asterisk (*) indicates a significant difference between the indicated groups, p < 0.001.       52 2.5 Discussion We have examined the distribution of RyR in adult rat ventricular myocytes. Our primary method was TEM because RyRs have a characteristic and unmistakable profile in well preserved and stained sections and because TEM provides direct visualization of the compartment in which the RyR are located.  The baseline for our measurements was established by examining junctions with t-tubules at the Z-line and junctions with the surface, Figure 2.1. These images displayed the expected components in the appropriate spatial configuration. First, the membranes of the sarcolemma and the SR are only about 10-15 nm apart. Second, the lumen of the SR is narrow and contains electron dense material that is thought to be largely calsequestrin. Third, the electron dense ryanodine receptors, or feet, are clearly seen on the SR membrane, spaced at regular intervals extending towards the t-tubule. The variable features are the size of the junction, which can only be determined from serial sections or tomograms, and the architecture; the SR forms a junction with a portion of the t-tubule, Figure 2.1A, or it can encircle most or all of the t-tubule, Figures 2.2E, 2.2F and 2.5A. All of the axial junctions displayed characteristics identical to those listed for junctions on t-tubules and on the surface (Figures 2.2A, 2.2B, 2.2C, 2.2D, 2.4, and 2.6).  The arrangement of RyRs in a native membrane is thought to be that of a regular lattice, with centre-to-centre distances between individual RyRs of 31.5 nm, or 44.5 nm on the diagonal (Yin et al., 2008). The distance I measured between RyRs in junctions with the t-tubule (44.3 ± 2.4 nm) and on axial junctions (40.0 ± 0.88 nm) is within this range. On the surface, I observed a roughly equal proportion of junctions that were doublets and singles. Comparable results have been seen in immunofluorescence images where RyR on the surface was distributed in doublets on either side of the Z-line (Chen-Izu et al., 2006).    53 We did not find cSR in ventricular myocytes, though it was readily visible in virtually all of the micrographs acquired from the atria, as expected (Dolber and Sommer, 1984; Jorgensen et al., 1985). The discrepancy between our findings and those reported in the literature likely stem from the different tissues that were examined; ventricular versus papillary myocytes. cSR in rat papillary muscle has RyR (Jorgensen et al., 1993), as well as CSQ (Jorgensen et al., 1988; Jorgensen et al., 1985) and the ability to store Ca2+ (Jorgensen et al., 1988). If the rat ventricular myocytes have cSR, it is very rare. These results indicate that there are structural differences between papillary and ventricular muscles that argue against using the former as a stand-in for the latter.  We searched extensively for RyRs in rat ventricular myocytes and throughout the SR but were able to find them in only three locations; in junctions with t-tubules on the Z-line, in junctions on the surface that may or may not straddle the Z-line, and in axial junctions running perpendicular to the Z-line. Axial tubules were readily visible and most of the profiles seen in sections, 75%, formed an axial junction with the adjacent SR. It is likely that the remaining 25% also formed junctions that were simply outside of the section and not visible. The length of the axial tubules was variable, but showed a peak at roughly one quarter of a sarcomere in length (mode = 0.26, Figure 2.3A). This was true of tubules both with, and without, junctions (Figures 2.3A and 2.3C). The fluorescence images, Figure 2.8, also show both short and long stretches of axial RyR and Cav1.2, so it is unlikely that short tubules are artifacts caused by sectioning.  Axial tubules have been observed using lipophilic membrane dyes, such as di-8-ANEPPS, in combination with optical microscopic techniques, but this approach only shows the presence of a tubule and cannot reveal the presence of a junction (Soeller and Cannell, 1999). Using electron   54 microscopic techniques, RyR have been observed adjacent to axial tubules (Tomita and Ferrans, 1987; Zhang et al., 2001). The significance of our results is in showing that most axial tubules in rat ventricle have junctions that are quite extensive, sometimes running all the way from one Z line to the next. They are also frequent, constituting 21% of all the junctions observed in thin sections. Given a median length of 510 nm and 8.5 rows of RyR, a single axial junction contains, on average, 109 RyR. Since t-tubule junctions contain between 120 and 260 RyR (Franzini-Armstrong, 1999; Soeller et al., 2007), I estimate that between 9% and 19% of the cell’s complement of RyR is deployed in axial junctions.  Most RyR are positioned in jSR opposite clusters of Cav1.2 in the apposing membrane, be it on the surface or in the TATS, but we and others have consistently found that there are RyR without adjacent Cav1.2, the extra-dyadic RyR. This is apparent in numerical analyses of immunofluorescence images that always demonstrate a significantly greater colocalization of Cav1.2 with RyR than vice versa (Mohler et al., 2005; Scriven et al., 2000a; Song et al., 2006). An example of an extra-dyadic cluster is shown in the image presented in Figure 2.8Aii. In contrast, every axial tubule looked like those displayed in Figures 2.8Aiii and iv, no extra-dyadic RyR are present. This visual impression was confirmed by the numerical analysis that showed a significantly greater colocalization of RyR with Cav1.2 in axial versus transverse tubules (Figure 2.8C). While we know that extra-dyadic RyR are adjacent to caveolin-3 and the signaling molecules housed therein, we do not yet know the function of the extra-dyadic RyR (Scriven et al., 2005).  Several lines of indirect evidence suggest that the axial junctions will be functional. First, its structural features are indistinguishable from those of the transverse junctions. The axial junctions have both RyR and CSQ, the distance between the axial tubule and the jSR membrane is   55 only 10-15 nm, and the axial tubule membrane anchors Cav1.2 opposite RyR in the jSR. Second, the structural features of the axial junctions in atria are identical to those of the ventricle (Figure 2.5B), and these atrial junctions are known to be active participants in ECC (Kirk et al., 2003). Third, the presence and frequency of RyR on the axial junctions is sufficient to explain some aspects of Ca2+ dynamics. Up to 20% of a rat ventricular myocyte’s spontaneous Ca2+ sparks originate from areas of the sarcomere that are too far from the Z-line to be attributed to t-tubule junctions (Lukyanenko et al., 2007; Shacklock et al., 1995). The large and relatively frequent assemblies of RyR in the axial junctions can easily account for this effect. In addition, it has been difficult to account for the comparable horizontal and longitudinal conduction velocities of Ca2+ waves if RyR are only on or near the Z-line (Subramanian et al., 2001). Again, my observations suggest that axial junctions could solve that problem. It is therefore reasonable to expect that axial junctions participate in normal excitation-contraction coupling as well as in the production of sparks and the spread of Ca2+ waves.  Tomography provided a striking view of the transverse and axial tubules, and their interconnections (Figures 2.6 and 2.7A movie sequence of the reconstructed 3D model, Figure 2.6C is available in the supplementary data) ( The most surprising aspect of the data is the tortured path the tubule follows, and the small diameter, 26 nm, of some of the tubules. Both aspects, small diameter and tortuosity, have been noted by others using EM techniques (Sommer and Waugh, 1976; Tomita and Ferrans, 1987), but not those using optical microscopy, possibly because the dyes used to label the tubules cannot diffuse through apertures as small as we have measured. This would preclude the finer structures from being observed using optical microscopes. Nevertheless, these techniques have revealed a remarkable increase in the number of   56 axial tubules in conjunction with a loss of transverse tubules in failing and chronically ischemic myocytes in the rat (Bito et al., 2008; He et al., 2001; Heinzel et al., 2008; Song et al., 2006).      57 3. The structure and functioning of the couplon in the mammalian cardiomyocyte1 3.1 Introduction The fidelity of information transfer across the plasmalemma is achieved, in part, by spatially confining second messenger networks to discrete subcellular domains. Each domain, which is often only nanometers in scale, contains the messenger’s targets as well as the molecular machinery needed to create the messenger. This architecture allows the parallel operation of multiple signaling pathways with little, or no, cross-talk even when composed of the same molecular components. Since a host of diseases are caused by breakdowns in these signaling networks, it is not surprising that mapping these domains, determining their molecular constituents and discovering how they function, develop and change over time, are critical goals for cell physiologists (Scott and Pawson, 2009). Excitation-contraction coupling (ECC) in cardiomyocytes, controlled by the second messenger Ca2+, is a perfect example of how information is spatially encoded in a nanodomain. A small influx of Ca2+ across the sarcolemma through Cav1.2 triggers a much larger Ca2+ release into the myoplasm through adjacent RyR2 located in the membrane of the jSR, a mechanism of ECC called Ca2+-induced Ca2+ release (CICR) (Fabiato, 1983, 1985). Cav1.2 and RyR are largely distributed in specialized cellular structures called couplons where the jSR and surface membranes are only about 15 nm apart. Couplon is a generic term applied to any apposition of jSR and surface membrane (either sarcolemma or t-tubule) bringing together functionally-linked                                                 1 This chapter has been published as a review, the data and hypothesis presented in section 3.2.2 provide the context for chapter 4.   58 clusters of Cav1.2 and RyR, and includes both dyads and triads (Rios and Stern, 1997; Stern et al., 1997).  Couplons, and the general cellular architecture of the cardiomyocyte, can be seen in the low power electron micrograph presented in Figure 3.1A. The thin actin-containing filaments are anchored at the Z-lines which repeat at regular distances of about 2 µm. Rows of mitochondria are aligned between the contractile filaments and the intercalated disk separates adjacent cells. Couplons can be seen on transverse tubules at the Z-lines, and the image contains numerous axial tubules, most of which are also likely to contain couplons (Asghari et al., 2009). An adult ventricular myocyte will have couplons distributed on the cell surface near the Z-lines in addition to those on transverse and axial tubules. Regardless of their location, they all share the same structural features, highlighted in Fig 1B, which are: 1) Close apposition of the t-tubule and SR membranes. 2) Electron dense material in the lumen of the SR, which includes the Ca2+ binding protein calsequestrin (CSQ). 3) Readily visible RyR2 monomers at seemingly regular intervals on the surface of the SR facing the t-tubule. Cav1.2 molecules cannot be distinguished from other intramembranous proteins in standard transmission electron micrographs.  The couplon’s architecture is essential not only for CICR, but also for insulating the cell’s many other Ca2+ dependent processes from the high local Ca2+ concentration required for RyR2 and Cav1.2 to function. Under normal physiological conditions the RyR2 have an affinity for Ca2+ in the µM range (Laver and Honen, 2008). If such a high concentration were attained within the myoplasm, it would activate numerous Ca2+ dependent processes, including proteolytic enzymes, resulting in many undesirable consequences. Spatially confining the proteins to the nanodomains defined by the couplons not only isolates CICR from the cell’s other Ca2+-dependent processes, it also reduces the number of Ca2+ ions that must enter the cell from the extracellular space. Recent   59 evidence indicates that the Cav1.2 clusters occupy a smaller membrane area than, and are positioned near the centre of, their adjacent RyR2 clusters (Scriven et al., 2010). This arrangement should further increase the probability of an entering Ca2+ ion finding its binding site on RyR2, further reducing the number of ions required for RyR2 activation.    Figure 3.1 Transmission electron micrographs of adult rat ventricular myocyte. A) Low magnification (7000x) of two myocytes. m – mitochondrion. ID – intercalated disk. Z – z lines. c – couplons. a – axial tubules. Scale bar = 2 mm. B) High magnification (120000x) of a single couplon on the Z-line. Scale bar = 100 nm. SR – sarcoplasmic reticulum. RyR2 – ryanodine receptor type 2. t – lumen of t-tubule. CSQ – calsequestrin.  Like most other voltage-gated channels, Cav1.2 inactivates as a function of time and membrane potential, but it is also inactivated by Ca2+ ions. Considerable evidence indicates that   60 these inactivating ions come primarily from the adjacent RyR2 (Adachi-Akahane et al., 1996). It is notable that Ca2+ released from other couplons is far less effective in promoting inactivation. This can be readily seen in genetically engineered mice in which the couplon architecture has been disrupted and the colocalization of Cav1.2 and RyR2 decreased. In these animals, many clusters of Cav1.2 are no longer adjacent to RyR2, and Ca2+-dependent inactivation of Cav1.2 is measurably delayed. Loss of rapid inactivation delays membrane repolarization, which prolongs the duration of the action potential (Chopra et al., 2009). These electrical perturbations are arrhythmogenic and potentially lethal.  Each couplon is physically separate from the others, being about 700 – 750 nm apart; an isolation that is also thought to be functional. Surprisingly, this architecture was predicted on theoretical grounds, through mathematical modeling, prior to its discovery (Stern, 1992), although empirical evidence rapidly followed. First came a series of observations describing Ca2+ ‘sparks’, which are stochastic openings of a few RyR2 that are localized neither producing Ca2+ sparks in other sites in the cell nor all-or-none Ca2+ transients (Cannell et al., 1994, 1995; Cheng et al., 1995; Cheng et al., 1993; Lopez-Lopez et al., 1994) despite releasing Ca2+ into a large volume. Second, results from electron microscopy and immunofluorescence clearly identified Cav1.2 arranged in clusters in the sarcolemma, directly opposite clusters of RyR2 in regions of the SR, junctional SR (jSR), where the two membrane systems are closely apposed (10-15 nm) in couplons (Carl et al., 1995; Sun et al., 1995).  The theoretical modeling, the sparks, and the structural data, all came together to provide the foundation for what is now considered the standard theory of cardiac ECC, local control theory. This theory states that under normal conditions intracellular Ca2+ release from the SR is through couplons that are spatially isolated from each other, and since a high local Ca2+ is required to activate CICR, the physical separation   61 produces a functional separation as well so that each couplon acts independently of the others. Under this paradigm the fundamental event in ECC is a Ca2+ spark, and an intact myocyte’s Ca2+ transient is the macroscopic sum of thousands of sparks that are synchronized by the action potential.  Local control theory has been immensely successful, but has some limitations. First, it treats each couplon as functionally identical. Recent findings suggest there are multiple types of couplons, in both atria and ventricle, that can be differentiated on the basis of their molecular partners (Schulson et al., 2011; Scriven et al., 2005). Many of the molecular partners are either directly involved in transmembrane Ca2+ transport or in modifying that transport, implying that different couplons function differently. This diversity should create a rich environment for controlling Ca2+ release and therefore the force of contraction. Second, local control theory does not explain how an individual couplon functions. Evidence indicates that only a small fraction of a couplon’s complement of RyR2 are activated with each action potential (Wang et al., 2004), but why this is so and what prevents the others from activating are both unclear. Recent evidence from our lab suggests a possible answer; not all RyR2 within a given couplon have the same sensitivity to Ca2+. These two hypotheses, multiple types of couplons and multiple Ca2+ sensitivities for RyR2 monomers within a couplon, along with their ramifications, and some of the obstacles to studying them, are the subject of this chapter.  3.2 Hypotheses 3.2.1 Couplons are diverse and some are variable amplifiers of ECC Although never explicitly stated, it has been thought that all couplons are similarly structured and therefore comparable in function. Recent evidence based on new, quantitative,   62 methods for analyzing cellular structure and molecular architecture has challenged these assumptions in both atria and ventricle.  Multiple types of couplons, distinguished by the molecules that colocalize with, and could modify the function of, RyR2, have also been identified in rat atria (Schulson et al., 2011). The majority of RyR2 clusters were associated with Cav1.2, suggesting that Cav1.2 mediated Ca2+ influx is the primary mechanism of ECC in this tissue. A statistically significant fraction, 10% - 15%, of those couplons were also associated with the Na+/Ca2+ exchanger, NCX, while a small proportion, 3% - 5%, were adjacent to caveolin-3. Atrial couplons differ in their probability of initiating a Ca2+ spark, with some having been designated as ‘eager’ sites, while others fail to spark (Mackenzie et al., 2001). The role of NCX is controversial, but where it is an integral part of the dyad, it could move Ca2+ into the dyadic cleft during the upstroke of the action potential leading to a ‘sensitized’ RyR2 array and an ‘eager’ Ca2+ spark site. So the differing architectures we’ve observed could explain these phenomena. Caveolin-3 anchors signaling molecules that affect virtually every protein involved in ECC and could readily augment or inhibit Ca2+ release.  The rat ventricular myocyte has been more extensively studied than the atrial. In ventricle each cell has roughly 30,000 clusters of RyR2 with an inter-cluster distance of approximately 550 nm (Scriven et al., 2010). We assume that all of these clusters are within couplons because an extensive TEM analysis found RyR2 distributed only in couplons, but nowhere else (Asghari et al., 2009). A small proportion of the clusters, 5% - 10%, colocalize exactly with NCX (Scriven et al., 2000a). The role of NCX in modifying Ca2+ levels within the ventricular couplon has been controversial. Empirical evidence from NCX null mice indicates that NCX contributes to Ca2+ influx during the action potential upstroke and sensitizes that couplon to subsequent Ca2+ influx via Cav1.2 (Larbig et al., 2010). NCX may also function to keep the intra-couplon Ca2+ levels low   63 during diastole so as to minimize Ca2+-dependent inactivation, and therefore maximize the availability of Cav1.2 during the next systole (Sher et al., 2008). Regardless of the role it plays, the presence of NCX within and near the couplon almost certainly modifies that couplon’s function. Two types of couplons have been distinguished by their size and the presence, or absence, of Cav1.2 (Scriven et al., 2010). The first were physically larger, contained about 75 RyR2 monomers and 10 individual Cav1.2 channels, and were more numerous; about 20,000/cell. The Cav1.2 were usually concentrated near the centre of these RyR2 clusters, which as previously noted, is an arrangement likely to ensure that Ca2+ ions permeating Cav1.2 have a high probability of binding to a nearby RyR2. The second group were less numerous, roughly 10,000/cell, and had only about 30 RyR2 monomers each. Importantly, these couplons were without Cav1.2; we refer to this group as the extra-dyadic RyR. Previous results from indirect immunofluorescence have demonstrated a population of RyR2 clusters without adjacent Cav1.2, but colocalized with caveolin-3 (Cav3). Since I cannot find RyR2 anywhere outside of a structure resembling a couplon, I have hypothesized that these smaller RyR2 clusters are adjacent to Cav-3. If true, there are at least two types of couplons in a ventricular myocyte, a Cav1.2 couplon and a Cav3 couplon, with a small percentage of the former containing NCX. The proposed architecture is diagrammed in Figure 3.2A.  The possibility of a Cav3 couplon is significant because of the numbers involved (30%-40% of the total) and the roles played by caveolin-3 in cells, particularly that of a signalosome; a place where signaling molecules are anchored and kept dormant by Cav3 until they’re activated by an appropriate stimulus. Two of the molecules anchored by Cav3, endothelial nitric oxide synthase (eNOS) and the β-AR, have been shown to directly impact ECC. eNOS is involved in the β-AR, muscarinic and frequency-dependent response of the heart (Champion et al., 2004;   64 Takimoto et al., 2005) while selective activation of the β-AR produces a potent positive inotropic response (Xiao et al., 1999). These data establish the localized effects of the signaling, and in combination with the structural data, suggest that the Cav3 associated RyR2 clusters can amplify CICR. This signaling microdomain would be a variable amplifier, with the extent of amplification determined by the level of activity of the adjacent signaling molecules. Interestingly, some of the other signaling molecules anchored by Cav3, such as PKA and PKC as well as eNOS, are also Ca2+-sensitive, strongly suggesting that these domains have a role in Ca2+ homeostasis and signaling. The Cav3 couplons are not randomly distributed, instead our data indicate that they are positioned significantly closer to the Cav1.2 couplons than the Cav1.2 couplons are to each other (Scriven et al., 2010). This would place them in an ideal location to respond to Ca2+ diffusing from the larger Cav1.2 couplons, and we know from studies of Ca2+ waves that a couplon’s nanospace is accessible to myoplasmic Ca2+ (Cheng et al., 1996). The smaller number of RyR2 within a Cav3 couplon, and its expected smaller release of Ca2+, would not prevent it from having a significant effect since the force-Ca2+ concentration relationship has a maximum slope of 4, so a small change in myoplasmic Ca2+ can have a dramatic effect on contractile force and cardiac output.   3.2.2 The RyR2 within a couplon form a disordered array How each of the couplons functions is, surprisingly, not understood. This is partly due to the fact that RyR2 are located on an interior membrane (the SR) and cannot be patch-clamped, which is the usual method for studying membrane-bound ion channels. In addition, RyR2’s enormous size, 104 exons and 4965 amino acids, makes it difficult to work with heterologous expression systems. Instead, its operation has been largely deduced from studying purified RyR2   65 in lipid bilayers and from the study of Ca2+ sparks, the latter being an indirect readout of RyR2 activity in intact cells. Two of the central and unresolved questions concern the number of RyR2 that open during a Ca2+ spark or a full-fledged Ca2+ transient, and how the positive feedback process of CICR is terminated. In other words, the mechanisms that control the RyR2 monomers aren’t fully understood.  During systole, do all of the RyR within a given dyad open, or just a few? The first idea, that all of the RyR in a couplon open, seems intuitive given the large Ca2+ influx into the confined space of the dyad. In addition, evidence indicates that RyR2 can exhibit coupled gating (Marx et al., 2000), which would be facilitated by a checkerboard orientation of the monomers allowing a positive allosteric interaction between them. This gives sound reason to believe that all RyR in a dyad would open. However, if they all open the single channel RyR2 current would have to be about an order of magnitude smaller (~0.05 pA/RyR2 given ~75 RyR/dyad) than is currently thought (Mejia-Alvarez et al., 1999). There is also strong experimental evidence that just a few RyR2 monomers open independently of the others (Wang et al., 2004). In sum, evidence does not support the ‘all RyR2 in a couplon open’ hypothesis. The second possibility is that only a few RyR2 in each dyad open during systole. The problem here is explaining why the remaining RyR2 don’t open given the high Ca2+ within the dyadic cleft. It has been postulated that the checkerboard organization of the RyR2 monomers provides strong allosteric inhibition that prevents most of them from opening (Stern, 1992), a feature of many mathematical models of dyadic function. However, there is no direct experimental evidence to support this. On the contrary, the existing evidence supports positive allosteric interaction (Marx et al., 1998; Marx et al., 2001b). Alternatively, or perhaps in addition, RyR2 monomers may have Ca2+-dependent inactivation. This was originally proposed by Fabiato   66 (Fabiato, 1985), but it would have to be ~200-fold stronger than previously thought to prevent most of the RyR2 from activating, particularly if there’s positive co-operativity (Wang et al., 2004). It was recently discovered that the dyads aren’t completely filled with RyR2, instead each is a ‘supercluster’ containing smaller RyR groupings, or archipelagos, with a number of single RyR2 present in each dyad (Baddeley et al., 2009) (Hayashi et al., 2009), Figure 3.2B. This has led to the idea that only a few of these archipelagos are activated during each systole, with the physical distance between them preventing runaway CICR within each dyad; in other words, local control theory on a microscale (Xie et al., 2010). However, since the distance between the archipelagos is small (<100 nm edge-to-edge; or about 3 RyR monomers wide (Baddeley et al., 2009), one would still expect all of them to be exposed to high Ca2+ and to activate.  RyR closure is hypothesized to be largely the result of Ca2+ depletion in the local SR. That idea arose from lipid bilayer experiments demonstrating that the myoplasmic sensitivity of RyR to Ca2+ was a function of luminal Ca2+ (Gyorke and Gyorke, 1998). This has been supported by single cell studies using low affinity Ca2+ indicators to directly monitor SR Ca2+, which clearly demonstrate a reduced, but highly localized, drop in SR Ca2+ (a Ca2+ blink), accompanying the Ca2+ sparks (Brochet et al., 2005). More recently, dual recordings of myoplasmic and SR Ca2+, performed simultaneously, confirm that Ca2+ sparks and Ca2+ blinks closely mirror one another in both space and time (Brochet et al., 2011). There is therefore considerable evidence to support luminal Ca2+ depletion in the local jSR as one factor controlling RyR excitability. However, the SR lumen can be refilled ~5 times faster than the spark amplitude recovers, so the sensitivity of the RyR2 recovers more slowly than does the SR Ca2+ content (Brochet et al., 2005).  Clearly, how a dyad is controlled remains an unsolved problem, preventing us from gaining a critical understanding of the basic processes of CICR and of RyR2 function. In addition,   67 RyR2 mutations cause human diseases, such as catecholaminergic polymorphic ventricular tachycardia and arrhythmogenic right ventricular dysplasia (Thomas et al., 2010), and both heart failure and hypertrophy involve changes in CICR (Gomez et al., 1997; McCall et al., 1998; Shorofsky et al., 1999). Therefore, understanding how RyR2s are controlled is essential for understanding both the normal and the diseased heart.  Electron micrographs of skeletal muscle triads show a fairly regular spacing between the RyR1 monomers, which suggested that they were distributed in a regular pattern. This was directly observed using freeze-fracture rotary shadowing of skeletal muscle RyR1, which clearly showed a checkerboard pattern of organization of RyR1 (Ferguson et al., 1984). Comparable rotary shadowing of cardiac muscle RyR2 hasn’t been performed. Instead, transmission electron micrographs of cardiac muscle show a fairly regular spacing between the RyR2 monomers, from which it has been deduced that they too are distributed in a regular array (Franzini-Armstrong, 1999; Protasi et al., 1997). More recent data indicates this may not always be the case.   68   Figure 3.2 Ultrastructure of a myocyte and a couplon. A) Hypothesized structure of an adult ventricular myocyte. Couplons are visible on the cell surface, and on both transverse and axial tubules. All couplons contain RyR2, which are opposite Cav1.2 in the larger couplons and Cav3 in smaller couplons. B) Current model for the disposition of RyR2 monomers within a couplon. An individual couplon from (A) is isolated, then rotated 90° to view individual RyR2 monomers and their disposition within a single couplon. The position of individual Cav1.2 molecules relative to RyR2 monomers is unknown.      69 Figure 3.3A shows a model of a tomogram that I acquired from a junction, located on the Z-line and within the interior of the cell. The viewer is looking straight at the junction en face; the RyR (Rock et al. 2005) and t-tubule membrane (green) are clearly visible. The widths of the RyR are in agreement with the expected value of about 30 nm. This dyadic cleft is clearly not filled with RyR, which agrees with previous observations (Baddeley et al., 2009; Hayashi et al., 2009). Instead, the RyR2 are grouped into smaller clusters, or archipelagos, including what appear to be isolated monomers; however neither the monomers nor the clusters appear to be distributed in a regular checkerboard.  To provide an unbiased estimate of RyR2 distribution, I created a 2D array of appropriately sized monomers in their expected checkerboard pattern (Figure 3.3B), and attempted to cover as much of the data as possible. The RyR2 array was allowed to both translate and rotate, but to account for the fact that the dyad isn’t filled, monomers that had less than 50% of their area overlapped by data were eliminated. The result, in Figure 3.3C, was poor, with ~60% of the data unmatched by RyR2. Clearly, a regular array does not fit this data. The next approach was to consider the data as a fractured array, where each section of the data was fitted with its own checkerboard of RyR2 monomers independently of the others. The result, shown in Figure 3.3D, is an improvement, but 37% of the data was still unmatched. Finally, I retained the concept of a fractured array, but allowed the RyR2 monomers to be organized both in the expected checkerboard array and in an unexpected edge-to-edge packing arrangement. This provided the best result and covered ~85% of the data (Figure 3.3E). These results imply that RyR2 monomers are in at least two different configurations within the dyad. Since allosteric interactions affect RyR2 function, there are at least two functional groups of RyR2 within a single couplon, but the functional significance of these differences has yet to be determined. Overall, based on this   70 observation I hypothesized that RyR2s within couplons are not distributed in a regular array and their relative positions are not fixed.    Figure 3.3 Tomogram of individual couplon from adult rat ventricular myocyte. A) Tomogram of a couplon on the Z-line. RyR2 monomers (Rock et al. 2005), jSR membrane (green). B) Array of RyR2 monomers interacting through subdomain 6. C) Best result of fitting the RyR2 array (B) onto the data (A) allowing the array to both rotate and translate. Individual RyR2 monomers had to overlap the data by 50% or they were removed from the array. The remaining RyR2 monomers covered only 40% of the data. D) The array was independently fitted to individual sections of the data, and covered 63% of it. E) As in D, but removing the requirement that individual RyR2 monomers could only contact their neighbours through subdomain 6. This covered 85% of the data. Scale bars are all 30 nm.    71 4. Non-uniform and variable arrangements of ryanodine receptors within mammalian ventricular couplons 4.1 Introduction The type 2 ryanodine receptor (RYR2) is an integral membrane protein of the cardiomyocyte sarcoplasmic reticulum (SR) that functions as a Ca2+-activated Ca2+ ion channel. Each receptor is a homotetramer, measuring roughly 29 nm x 29 nm x 12 nm, which can be readily identified in electron micrographs based on its location within the dyadic cleft and on its size and shape (Carl et al., 1995; Sun et al., 1995). Rotary shadowing studies of type 1 ryanodine receptors (RYR1) in skeletal muscle triads (Ferguson et al., 1984) and numerous transmission electron micrographs of cardiac muscle (Franzini-Armstrong, 1999) left the impression that the tetramers filled the dyadic cleft, forming a defect-free crystalline array, often referred to as a ‘checkerboard’. The array’s formation is thought to be an intrinsic property of the protein that would reflect the homotetramer’s four-fold symmetry, whereby adjacent tetramers were non-covalently connected through their adjacent clamp domains (Liu et al., 2004). This is also thought to provide the structural basis for inter-protein allosteric interactions (Marx et al., 2001b; Porta et al., 2012b). Later on, electron tomography and super-resolution fluorescence microscopy revealed that the dyad contained sub-arrays that did not completely fill the cleft, although neither technique had the resolution to determine the position and orientation of individual tetramers (Baddeley et al., 2009; Hayashi et al., 2009). A single-tilt tomogram with higher resolution indicated that the sub-arrays were unlikely to be fitted with a simple checkerboard (Asghari et al., 2012). RYR1 tetramers, purified from skeletal muscle and inserted in artificial bilayers, spontaneously formed two different types of array that depended on the free Mg2+ concentration.   72 Using a nominally Mg2+ free buffer, the tetramers formed a checkerboard, but with the addition of 4 mM Mg2+ the tetramers were more densely packed in a side-by-side orientation although there was no physical contact between them (Yin et al., 2008; Yin et al., 2005b). The organization of the tetramers at the expected intracellular free Mg2+ concentration of ~1mM was not investigated. Whether RYR2 behaves similarly, and if such changes can occur in vivo, is unknown. In this chapter, I examined dual-tilt tomograms to directly visualize the position of individual RYR2 tetramers in adult rat ventricular myocytes. When fixed in situ, where the Mg2+ is ~1 mM (Tashiro et al., 2009; Tursun et al., 2005), en face views of the dyads showed RYR2 in a variety of patterns and orientations that were neither uniform nor regular. I obtained the same results from cells that were fixed after enzymatic dissociation or from cells that were fixed, permeabilized with the free Mg2+ set to 1.0 mM, as well as from sections of juvenile human ventricle. The tetramer distributions could be moved into more regular arrays by lowering the free Mg2+ concentration to 0.1 mM, or by phosphorylation, both of which resulted in a largely checkerboard arrangement, whereas high Mg2+ (4 mM) produced a more densely packed configuration where the tetramers were largely side-by-side. Changes in tetramer positioning were visible at the earliest time point I examined, which was ten minutes.  I conclude that the positioning of RYR2 tetramers within mammalian dyads is dynamic, that individual tetramers can both rotate and translate independently of each other, and that they do so relatively quickly in response to a change in local factors.   4.2 Materials and methods The experiments used ventricular myocytes from adult rats as well as left ventricular myocytes from humans. Animal handling was done in accordance with the guidelines of the   73 Canadian Council on Animal Care and approved by the animal research committee of the University of British Columbia (UBC). Human tissue was acquired from informed subjects and the study was approved by BC Children’s Hospital Research Ethics Board. All chemicals were purchased from Sigma-Aldrich (Oakville, ON) unless otherwise stated.  4.2.1 Experimental protocol Isolation of rat ventricular myocytes was performed as previously described (Rodrigues and Severson, 1997). In the case of human tissue, small sections of the left ventricle were obtained from patients undergoing coronary artery or valve replacement surgery. Within a minute of excision the sections were cut into cubes roughly 1 mm on a side then immersed in fixative (4% paraformaldehyde and 2.5% glutaraldehyde). The tissue blocks were then post-fixed, dehydrated, embedded in resin and stained for electron microscopic and tomographic analyses (Asghari et al., 2009).   4.2.2 Identification and placement of ryanodine receptors Dyadic clefts, regardless of their intracellular location, are complex three-dimensional structures whose juxtaposed membranes are seldom, if ever, parallel. Even when they appear to be so, undulations in the membranes are commonplace and en face images of the jSR, acquired from a vantage point within the cleft, invariably include elements of membrane that intersect the plane of view. I therefore viewed the tomograms using Amira (VSG, Burlington, MA) in all three dimensions to enable a positive identification of each structure. The en face views obtained from the 3-dimensional tomogram were converted to a stack of TIFF images spaced 1 nm apart and passing through the entire width of the dyadic cleft. The stack was then read into RyR-fit (written   74 by D. Scriven) which allowed me to position a square, 29 nm on a side and outlined in red, over each RYR2 that had been identified; 29 nm is at the upper end of the range of sizes reported for the tetramers’ myoplasmic domain (Radermacher et al., 1994; Samso et al., 2005; Yin et al., 2005b). Placement of the tetramer, to 0.5 nm, was by eye and the orientation could be adjusted in 1° increments. Nearest-neighbor distances were calculated from the tetramers’ centers. This approach allowed me to compensate for any curvature of the cleft since receptor clusters could be accurately visualized in whatever plane they were in focus. In addition, rocking the image back and forth through one or two planes of a tetramers’ plane of focus enabled me to position them precisely.   4.2.3 Statistical analyses Data were reported as mean ± S.D., and significance was evaluated by non-parametric Kruskal-Wallis test, while pairs of data were analyzed using the Mann-Whitney test with values of p<0.05 being considered significant.  4.2.4 Supplemental method  I used male Wistar rats, 200 – 300 grams, (Charles River Laboratories, Wilmington, MA) that were given an intraperitoneal injection of 2 ml of 1000 units of heparin (Hepalean; Organon, Mississauga, ON) followed 30 minutes later with an intraperitoneal injection of 2 ml of sodium pentobarbital (240 mg/ml; MTC Pharmaceuticals, Cambridge, ON). The experiments proceeded only after hard pressure on the footpad failed to produce a withdrawal reflex. The isolation technique is based on the method of Rodrigues and Severson: The hearts were excised, hung on a Langendorff apparatus and perfused for 5 min. at 37°C with a nominally   75 Ca2+-free physiological saline solution (PSS) (in mM): 138 NaCl, 5 KCl, 0.3 KH2PO4, 0.3 Na2HPO4, 10 HEPES, 15 D-glucose, 1 creatine, 1 carnitine, pH 7.4) that had been equilibrated with 95% O2/5% CO2. Cell dissociation was initiated by switching to a perfusate of PSS containing 0.5 mg/ml Type II Collagenase (Worthington Biochemical, Lakewood, NJ) and 1 mg/ml bovine serum albumin. When the heart began to soften, the ventricles were cut free and sliced into small chunks, which were gently shaken to dislodge cells. These were filtered through a 200 µm nylon mesh (Nitex) into fresh PSS. There were typically greater than 90% quiescent, rod-shaped, cells. Calcium sparks in permeabilized myocytes Isolated cells were suspended in a solution (final volume 50 µl) containing (mM): potassium aspartate 100; KCl 20; EGTA 0.5; MgCl2 0.75; and HEPES 10; (adjusted to pH 7.2 with KOH), and placed in the experimental chamber for 15 min. Laminin-coated (5 µg/cm2; Roche Diagnostics, Laval, QC) glass-bottom grid-50 µ-dishes (ibidi, Verona, WI) were used as the experimental chamber to relocate cells for electron microscopic examination later. The cell membrane was permeabilized by adding 0.005 % (w/v) saponin for 30 s. After 30 s the bath solution was exchanged to a saponin-free internal solution composed of (mM): potassium aspartate 100; KCl 15; KH2PO4 5; NaATP 5; EGTA 0.5; CaCl2 0.2; phosphocreatine 10; HEPES 10; fluo-4 potassium salt 0.03 (Molecular Probes, Eugene, OR); creatine phosphokinase 5 U ml−1; 40000 MW dextran 8 %; pH 7.2 (adjusted with KOH). In addition, the solution contained MgCl2 at one of the following concentrations (in mM): 2.4, 5.5 or 8.0, giving a free [Mg2+] in solution of 0.1, 1, or 4 mM respectively. In all cases the free [Ca2+] was 100 nmol/l. The concentrations of MgCl2 and CaCl2 were determined using WEBMAXC Extended   76 ( with a temperature of 22 ̊C, 0.1N ionic strength and pH of 7.2. After a 10 min incubation period Ca2+ sparks were recorded using a Zeiss AxioObserver inverted microscope with a 20X/1.4 NA water immersion objective and a narrow bandpass filter (Semrock, Rochester, NY) optimized for fluo-4. Images were captured on an Evolve 512 CCD camera (Photometrics, Langley, BC) and the sparks were analyzed using Fiji and spark frequencies were expressed as sparks/s/mm2. The SR Ca2+ content was evaluated by the addition of 20 mM caffeine. The cells were immediately fixed in a solution containing 4% paraformaldehyde and 2.5% glutaraldehyde for 2 hours at room temperature and then prepared for electron- microscopy (EM) as described below. Phosphorylation After baseline Ca2+ sparks were recorded for 60 s in permeabilized myocytes exposed to a free Mg2+ concentration of 1 mM, and a free Ca2+ concentration of 100 nmol/L, a saponin-free internal solution containing 1 µmol/l thapsigargin, was added to the myocyte suspension to prevent Ca2+ uptake into the SR. After incubating for 10 min the cells were washed two times with the saponin-free internal solution, after which the cells were bathed with internal solution to which I added a phosphorylation cocktail consisting of (µmol/l): 10 c-AMP, 10 3-isobutyl-1-methylxanthine, 10 okadaic acid, 0.5 calyculin A and 1 thapsigargin, pH 7.2 (KOH). This cocktail activates kinases including protein kinase A and Ca2+-calmodulin-dependent protein kinase II; it deactivates phosphodiesterases and inhibits phosphatases PP1 and PP2. After a 10 min incubation period Ca2+ sparks were recorded for 60 seconds after which 20 mM caffeine was applied, then the cells were fixed in a solution containing 4% paraformaldehyde and 2.5% glutaraldehyde for 2 hours at room temperature and prepared for EM. The addition of thapsigargin in this protocol   77 prevented the Ca2+ content of the SR from rising, which after the addition of caffeine, just before fixation, was lower than in the control cells (1 mM Mg2+) (Figure 4.6) due to the accelerated spark rate (Figure 4.6A). The cells for the phosphorylation, isolated fixed, and 1 mM Mg2+ protocols were obtained at the same time from each animal, then split into these three groups and processed accordingly. The results obtained from the isolated fixed, and the 1 mM Mg2+ experiments (Tables 4.1 and 4.2) served as controls for the phosphorylation. Confirmation of RYR2 phosphorylation We used a standard Western blot protocol to confirm phosphorylation of RyR2 by the phosphorylation cocktail. The 6X sample buffer included: 350 mM Tris-Cl (pH 6.8), 30% glycerol, 10% SDS, 600 mM dithiothreitol, NaF (6 mM) and a protease inhibitor (P-8340, Sigma) was directly added to the cells before and after treatment. The samples were frozen in liquid N2 for storage at -80 ̊C until use. Protein (quantified via a Bradford protein assay, Bio-Rad 500-0001, Hercules, CA) was separated on 6% SDS- polyacrylamide gels and transferred to nitrocellulose membranes overnight at 4oC (BioRad, Hercules, CA). A polyclonal antibody to the Phospho Serine-2814 (A010-31 AP; Badrilla Ltd., Leeds, UK) was used to detect whether RyR2 was phosphorylated. To establish the loading control, each membrane probed with phospho-antibody was washed, duplicates were split and one was reprobed with an antibody to calsequestrin (rabbit polyclonal; PA1-913, Affinity BioReagents) while the other was reprobed with an antibody to caveolin-3 (mouse monoclonal; 610420, BD Biosciences). Protein bands were visualized using the SuperSignal West Femto Chemiluminescent Substrate kit (Thermo Scientific, Rockford, IL).    78 In situ fixation In rats, after the foot pad reflex failed to produce a response, the chest was opened, the aorta and vena cava were cannulated and the hearts were perfused for 10 min with physiological saline solution (PSS) followed by a fixative containing 4% paraformaldehyde, 2.5% glutaraldehyde and 4 mM CaCl2 in a 0.1 mol/l cacodylate buffer (pH 7.4; Canemco & Marivac Inc, Lakefield, PQ), for 10 min. The left ventricle was then removed, cut into small blocks and the samples immersed in fixative for about 2 hours, after which I proceeded as described below. Preparation for tomography Tissue or cells immersed in fixative were cyclically microwaved (2 min on, 2 min off, 2 min on) in a vacuum using a Pelco 3450 laboratory microwave (Ted Pella Inc., Redding, CA), at power 5, then rinsed and microwaved twice in 0.1 mM cacodylate buffer for 40 seconds at power 1, then post-fixed with 1% OsO4 solution (EMS, Hatfield, PA), at power 1, then cyclically microwaved twice. En bloc staining of samples was done with 2% aqueous uranyl acetate (Ted Pella Inc.), cyclically microwaved twice at power 1, then rinsed three times with distilled water followed by dehydration in ethanol (50-100% in steps of 10%; microwaved 1 min at each dilution on power 3), then embedded in a mixture of Epon and Spurr’s resin (Asghari et al., 2009). Image acquisition and tomography Grids were placed in a rotating, dual high-tilt stage and observed in the 200 kV Tecnai G2 Sphera transmission electron microscope (FEI, Hillsboro, OR). A suitable junction was imaged twice from orthogonal directions with serial tilt views ranging from +65° to -65° at 1° increments using TIA (FEI), an automated acquisition program. The 3D volumes were reconstructed using   79 either real-space back-weighted projection or the simultaneous iterative reconstruction technique (SIRT) implemented in Inspect 3D (FEI). Dual-tilt alignment and visualization was done using Amira 5.3 (VSG, Burlington, MA). Dual data sets examined using the Multi Planner Viewer, a sub-application in Amira 5.3. No image processing steps beyond the contrast stretch were employed.  4.3 Results The image displayed in Figure 4.1Ai is a single plane extracted from the dual-tilt tomogram of a rat myocyte fixed in situ, and shows a triad with characteristic jSR (arrows) and its ryanodine receptors on either side of a t-tubule (double arrow). Viewpoints within the volume of the tomogram are determined by the position of orthogonal planes which are outlined in different colors; XY in red, YZ in green and XZ in blue. In Figure 4.1Ai the XZ plane has been positioned to parallel, as nearly as possible, the jSR membrane, but to be within the cleft and to bisect the ryanodine receptors on that side of the triad. The intersection point of all three planes has been positioned within a single ryanodine receptor identifiable by its characteristic shape (roughly square) and size (~29 nm on each side), which is visible in all three orthogonal views; XY (Figure 4.1Ai), YZ (Figure 4.1Aii) and XZ (Figure 4.1Aiii). The XY and YZ views demonstrate that the intersection point of the planes is not within the jSR or the t-tubule membranes. A second YZ plane (yellow line in Figures 4.1Ai, iii) is within the t-tubule membrane, which in the en face view (Figure 4.1Aiii) has a similar size and shape as a RYR2. We used this procedure to differentiate the ryanodine receptors from membranes and other structures in this and subsequent tomograms. Additional examples of identifying membrane and RYR2 are in Figure 4.2, and the complete   80 tomogram of this junction, in an XZ orientation, can be viewed in supplemental movie M1 (   81     82 Figure 4.1 RYR2 distribution in a rat cardiomyocyte fixed in situ.  Images are from a tomographic study of a 200 nm thick section of left ventricular tissue. (A) Three orthogonal planes intersecting at a point within the image. Each plane is associated with a color: XY – red; YZ - green and XZ – blue; scale bars are 60 nm. (i) A slice through the XY plane of the tomogram showing a triad in which the YZ (green line) and XZ (blue line) planes intersect in the middle of a ryanodine receptor. A second YZ plane (yellow) intersects the XZ plane in the t-tubule membrane; single arrows – jSR, double arrow – t-tubule. (ii) A slice along the YZ plane of the data set at the level of the green line in Ai. (iii) A slice along the XZ plane (blue line in Ai & ii). (B-D) A single 1 nm-thick slice from the XZ plane of the tomogram in various stages of the process used to identify the position and orientation of the tetramers. The scale bars are all 30 nm. (B) (i) A single XZ slice from the tomogram. (ii) The same slice with the tetramers identified by red circles, 41 nm diameter. (C) Determining the position and orientation of the tetramers – the yellow square in (i) is the region of interest (ROI). (ii) The ROI magnified 15%, showing four tetramers. (iii) Outlines of the tetramers drawn by hand. (iv) The dotted outlines with accurately sized (29 nm) RyR2 tetramers (red boxes) manually positioned over them. (v) Final position and orientation of the tetramers. (D) (i) The distribution and orientation of all of the tetramers in the couplon positioned over the image. (ii) The tetramers and their center-to-center nearest-neighbor distances (nm).    83 An enlarged en face view of the junction is presented in Figure 4.1Bi, and in Figure 4.1Bii red circles with diameters of 41 nm (equivalent to the diagonal of a 29 nm square tetramer) centered over the areas identified as RYR2. Areas that are stained, but were not identified as RYR2 tetramers are sections of either SR or t-tubule membrane that were within the plane of view. It is apparent from these images that the RYR2 are not distributed in a well-ordered checkerboard array, an observation that agrees with our previously published single-tilt tomogram (Asghari et al., 2012). The increased clarity and resolution of a dual-tilt tomogram (Mastronarde, 1997) enabled me to estimate the position and orientation of each tetramer as depicted in Figure 4.1C. Individual receptors were first outlined with a dashed yellow line (Figures 4.1Ci, ii and iii), then fitted with squares (red), 29 nm on a side, Figures 4.1Civ and v. The result was an en face view of the junction in which the position and orientation of the ryanodine receptors were identified (Figure 4.1Di) and enabled the nearest-neighbor center-to-center distances (NND) to be calculated (Figure 4.1Dii). I acquired 11 tomograms from 6 hearts fixed in situ and examined 215 tetramers within dyads located on the cell surface as well as on both axial and transverse tubules. I also measured the NND of 56 tetramers (3 tomograms) acquired from myocytes that were enzymatically dissociated then fixed (Figure 4.3B), and a further 30 tetramers (2 tomograms) from dissociated myocytes whose membrane was permeabilized with saponin and the cell was incubated in a solution containing 100 nmol/L Ca2+ and 1 mM Mg2+ prior to being fixed. The histogram of the combined 301 tetramers’ NND, Figure 4.3A, shows a broad and bimodal distribution with modes at 32 and 38 nm. Separate histograms for each of the data sets are available in Figure 4.4. Although the tetramers’ positions and orientations were not uniform, an individual tetramer’s position relative to its neighbors could be broadly classified using a set of criteria   84 derived from the bimodal distribution of their NND and from visual examination of the images. I considered a tetramer to be in a checkerboard configuration relative to its neighbor(s) if their sides were parallel and separated by ≤3 nm, and overlapped by ≤19 nm (2/3 of its length). If those criteria were fulfilled but the overlap exceeded 19 nm the tetramers were considered to be side-by-side. Some tetramers had neighbors in both configurations while others had none and were considered isolated. These criteria accommodate the wide range over which neighboring tetramers can overlap, which, as is evident from the images, can vary from complete to just touching at the corners. The NND, sorted using the above criteria, are redisplayed in Figure 4.3B. The arrows show the NND for side-by-side (single arrow) and checkerboard configurations (double arrow) based on the values determined for purified RYR1 in vitro (Yin et al., 2005b) and assuming a 29 nm sized tetramer. Of the 301 tetramers that were identified (15 tomograms), 140 (46.5%) were in a checkerboard orientation, 117 (38.9%) were side-by-side, 24 (8.0%) were isolated and 20 (6.6%) had neighbors in both configurations (Table 4.1). The mean nearest neighbour distances of the tetramers in a checkerboard orientation was 36.9 nm +/- 2.2 nm (median 37.0 nm), while those in a side-by-side configuration was 30.7 nm +/- 1.2 nm (median = 30.7 nm), and those that were isolated was 42.1 nm +/- 9.3 nm (median = 40.3 nm) (Table 4.2).     85   Figure 4.2 Two different sets of orthogonal views of the tomogram presented in Figure 4.1.  Fig. 4.2 Ai is in the XY orientation and shows that the XZ (blue) and YZ (green) planes intersect within the jSR membrane; the jSR is indicated by the white arrow. This is also apparent in the YZ orientation (Fig 4.2 Aii) showing the intersection point of the XY (red) and XZ (blue) planes. In the en face orientation, Fig. 4.2 Aiii, it is not possible to identify that point as a RYR2 or a section of membrane without the orthogonal views in Ai and Aii. B shows a comparable set of images where the intersection point of the planes is now centered within a RYR2, and well above the jSR membrane.      86    Figure 4.3 (A) Histogram of the combined center-to-center NND of three groups of rat cardiomyocytes, namely: fixed in situ; isolated and fixed; isolated, permeabilized, the Ca2+ and Mg2+ concentrations set to 100 nmol/L and 1 mM respectively, and then fixed. (B) The NND in (A) were sorted into the indicated categories and redisplayed. The NND for purified RYR1 tetramers, calculated using the formulation of Yin (Yin et al., 2005) but assuming a 29 nm square tetramer, are indicated for the side-by-side (single arrow) and checkerboard (double arrow) configurations.    87    Figure 4.4 Displays histograms of the tetramer’s nearest neighbor distances.  (A) NND of the myocytes fixed in situ. (B) NND of the myocytes enzymatically isolated and then fixed. (C) myocytes that were enzymatically isolated, permeabilized and incubated for 10 minutes in a solution with free Ca2+ and Mg2+ concentrations of 100 nmol/L and 1 mM respectively  and then fixed.   88   Table 4.1    Tetramer arrangement Quantitative analysis of the tomographic data indicating number of the tetramers studied and number of the dual tilt tomograms used in each condition.     89    Table 4.2   Nearest neighbor centre-to-centre distances (nm)      90 To determine whether the RYR2 distribution in human dyads was comparable, I obtained left ventricular tissue from patients undergoing cardiac surgery to repair valve or coronary artery defects; none of the patients were in heart failure or displayed any evidence of hypertrophy. The tomogram of a dyad, from the left ventricle of a 9 year old male, and the analysis of its RYR2 distributions is displayed in Figure 4.5. Orthogonal views are displayed in Figure 4.5A, with the intersecting planes positioned over a single ryanodine receptor. The mitochondrial cristae are clearly visible and demonstrate that the cell was well preserved. It is apparent from Figures 4.5Ai and 4.5Aii that the distance between the t-tubule and jSR membranes is variable, with the result that a single XZ plane (blue line Figure 4.5Ai) cannot transect all of the visible ryanodine receptors. In these cases multiple XZ planes were needed to determine the position and orientation of the receptors and in this instance I display two, separated by 8 nm. The en face views of those planes are displayed in Figures 4.5Bi and ii, and the ryanodine receptors are identified by the 41 nm diameter red circles in Figures 4.5Ci and ii. The fitted receptors are shown in the en face images in Figures 4.5Di and ii, and their summed distribution is in Figure 4.5Ei. Of the five human dyads examined, I identified 84 RYR2 tetramers, 35 (41.7%) of which had neighboring tetramers in a checkerboard configuration, 38 (45.2%) were side-by-side, 5 (6.0%) had a neighboring tetramers in both configurations and 6 (7.1%) of the tetramers were isolated (Table 4.1). The mean NND of those with a checkerboard configuration was 37.2 nm ± 1.8 nm (median 37.2) while the mean NND of the tetramers considered side-by-side was 30.2 ± 1 nm (median 30.2 nm) (Table 4.2). A histogram of the NND is displayed in Figure 4.5Eii. The results are virtually identical to those obtained from rat ventricular myocytes (Figures 4.1 and 4.3).    91       92 Figure 4.5  RYR2 distribution in a human left ventricular myocyte.  A) Orthogonal views, (i) XY, (ii) YZ, (iii) XZ, 1 nm thick, through a dyad, with the intersecting planes positioned over a single RYR2. Double arrow, t-tubule; single arrow, SR. Scale bars are 60 nm. B) Two XZ images separated in Y by 8 nm. Scale bars (B-E) are 30 nm. C) B with the tetramers identified by a red circle 41 nm in diameter. D) Final position and orientation of the tetramers. E) (i) The image displays all of the tetramers along with their nearest-neighbor distances (nm). (ii) Histogram showing the NND of all 84 tetramers identified from the dyads of 5 patients.    93 I hypothesized that the bimodal NND could be explained, in part, by the cells’ expected free Mg2+ concentration, 1 mM, which is between those used to produce the side-by-side and checkerboard arrays of purified RYR1 in vitro. We therefore permeabilized isolated rat myocytes with saponin and incubated them for ten minutes with solutions containing a free Mg2+ concentration of either 0.1 mM or 4 mM while the free Ca2+ concentration was 100 nmol/L. Ca2+ spark frequency and caffeine transients were recorded prior to fixing the myocytes in place, on the coverslips, for electron tomography. Changing the free Mg2+ concentration had the expected effects on the frequency of Ca2+ sparks, which was significantly increased by 0.1 mM Mg2+, and significantly decreased by 4.0 mM Mg2+ (Gusev and Niggli, 2008). The caffeine transients were not significantly affected by the different free Mg2+ concentrations (Figure 4.6).  The en face images acquired from cells incubated in 0.1 mM Mg2+ (Figure 4.7) are remarkable in that the RYR2 are almost all in the checkerboard arrangement. This pattern was obvious even in the raw data (Fig 4.7Aiii) before the tetramers were positioned, (Figure 4.7B), and their NND calculated (Figure 4.7C) and was a consistent observation for the 78 tetramers (5 tomograms, 5 cells, 4 rats) that I examined. Of those tetramers, 64 (82.1%) were in a checkerboard configuration with a mean NND of 37.2 nm ± 1.8 nm, while only 4 (5.1%) were side-by-side with a mean NND of 31.0 nm ± 0.4 nm (median 31.0 nm). Nine (11.5%) of the tetramers were isolated and 1 (1.3%) had neighbors in both configurations. The configurations and their NND are listed in Table 4.1 and a histogram of the combined NND is displayed in Figure 4.7C. Additional images from this and subsequent tomograms as well as movie sequences through their en face views are available in Figures 4.8, 4.9 and 4.10 and supplemental movies M2-M4 (   94  Figure 4.6   Ca2+ sparks frequency and SR content.  The spark frequency (A) and the SR content (B) for cells that were permeabilized and exposed to the indicated solution plus 100 nmol/L free Ca2+ for 10 minutes prior to recording. Incubating the cells for ten minutes in the phosphorylation cocktail or in 0.1 mM Mg2+ both produced significant increases in spark frequency compared to 1 mM Mg2+. The 4.0 mM Mg2+ produced a significant decrease and the different solutions produced no significant differences in SR Ca2+ content.. The phosphorylation cocktail ensured that the SR Ca2+ content was roughly comparable between the different treatments, and would have activated PKA, Epac and CamKII.  (I/Io) normalized Intensity.   A B 0 10 20 30 40 50 60 70 80 90 Phosphorylation cocktail 1 0.1 4 sparks/s/mm2   1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8 Phosphorylation cocktail 1 0.1 4  I/Io mmol/l free Mg2+ mmol/l free Mg2+   95  Figure 4.7 RYR2 distribution in a permeabilized rat cardiomyocyte with a free Mg2+ concentration of 0.1 mM.  (A) An XZ image of the dyadic cleft. (B) Position and orientation of the tetramers in A. (C) The tetramers are displayed along with their NND. Scale bars are 30 nm. (D) Histogram showing the NND of all between 77 tetramers identified in 5 similarly treated cells.   96    Figure 4.8 Additional images of the tomograms for cells that were permeabilized and exposed to 0.1 mM Mg2+.     97   Figure 4.9 Additional images of the tomograms for cells that were permeabilized and exposed to 4 mM Mg2+.    98     99 Figure 4.10 Additional images of the tomograms for cells that were permeabilized and exposed to cAMP with phosphodiesterase and phosphatase inhibitors. Figures 4.8, 4.9 and 4.10 provide additional images of the tomograms for cells that were permeabilized and exposed to 100 nmol/L Ca2+ and to one of: (4.8) 0.1 mM Mg2+; (4.9) 4 mM Mg2+; (4.10) cAMP with phosphodiesterase and phosphatase inhibitors. In each of the figures (A) shows the tomogram in XY (i), YZ (ii) and XZ (iii) orientations with the planes intersecting within an RYR2 tetramer. Single arrow – jSR; double arrow – t- tubule. (B) (odd numbers) displays an XZ view, or multiple XZ views separated in Y by the indicated values (6 nm between 4.9i and 4.9iii; 6 nm between 4.10i and iii; 20 nm between 4.10i and v), and (even numbers) highlights the RYR2 with a red circle 41 nm diameter. (C) (i) shows magnified views of one of the XZ planes where the insets highlight several RYR2 (ii-v). The magnified RYR2 are outlined by hand and fitted with appropriately positioned and oriented tetramers. We noted that the SR was enlarged when incubating the cells in 0.1mM Mg2+, but as Fig 4.8D demonstrates, the mitochondrial (M) cristae are well preserved indicating that the effect is not due to osmotic stress. The cause of this effect is unknown. T, t-tubule; SR, sarcoplasmic reticulum.               100 In contrast, images acquired from cells incubated in 4.0 mM Mg2+ produced a very different result. Two en face images of a dyadic cleft (Figure 4.11Ai, Bi), separated by 6 nm, demonstrate that the tetramers were more densely packed (Figure 4.11Aii, Bii). The NND for this cell are displayed in Figure 4.11Ci, and comparable results were obtained from the 5 tomograms I examined (5 cells, 5 rats). In these tomograms I identified 97 tetramers, 65 (67.0%) were side-by-side with a mean NND of 29.4 nm ± 1.0 nm (median 24.0 nm), and 18 (18.6%) were in a checkerboard configuration with a mean NND of 34.7 nm ± 3.0 nm (median 34.3 nm). The results are listed in Table 4.1 and a histogram of their combined NND is displayed in Fig 4.11 Cii. We then assessed the effect of phosphorylation on tetramer distribution in permeabilized myocytes in the presence of 1 mM free Mg2+ and 100 nmol/L free Ca2+. This increased the Ca2+ spark frequency (Figure 4.6), and a Western blot demonstrated that the receptor had been phosphorylated on S2814 (Figure 4.12A, B). The change in phosphorylation status and function were paralleled by changes in the tetramer distribution, as shown in Figure 4.13. Three en face planes of the tomogram are displayed both as raw data (Figure 4.13Ai, Bi, Ci) and with their tetramers positioned (Figure 4.13Aii, Bii, Cii). Figure 4.13Di displays the distribution and NND of these tetramers. The 56 tetramers identified in 5 different tomograms demonstrated that 50 (89.3%) were in a checkerboard orientation with a mean NND of 37.3 nm ± 1.8 nm, with only 3 (5.4%) that were side-by-side. The data are listed in Table 4.1 and a histogram of their NND is shown in Figure 4.13Dii.     101  Figure 4.11 RYR2 distribution in a permeabilized rat cardiomyocyte with a free Mg2+ concentration of 4.0 mM. Scale bars are 30 nm.  (A) (i) XZ image of the dyadic cleft. (ii) Final position and orientation of the tetramers that were well defined in Ai. (B) (i) XZ image of the dyadic cleft 6 nm deeper in Y. (ii) Final position and orientation of the tetramers that were well defined in Bi. (C) (i) The distribution and orientation of all of the tetramers along with their nearest-neighbor distances in nm. (ii) Histogram showing the NND of all 97 tetramers identified in 5 similarly treated cells.   102      103 Figure 4.12 Western blots. (A) An average of three Western blots demonstrated that the phosphorylation cocktail produced a significant increase in S2814 phosphorylation, an epitope phosphorylated by both CaMKII and PKA. (B) displays Western blots of the proteins selected as controls, calsequestrin and caveolin-3, and the normalized intensity of the bands. (C) An average of three Western blots demonstrated that the phosphorylation cocktail does not produce a significant increase in S2808 phosphorylation, another epitope phosphorylated by both CaMKII and PKA.     104  Figure 4.13 RYR2 distribution in a permeabilized rat cardiomyocyte dyad after phosphorylation. Scale bars are 30 nm. Because of the unevenness and curvature of the dyad three separate planes A, B (6 nm deeper in in Y than A) and C (20 nm deeper in Y than A) were required to identify all of the tetramers. Ai, Bi, Ci are raw images, Aii, Bii and Cii are the final orientation and position of the tetramers. (D) (i) The distribution and orientation of all of the tetramers along with their NND in nm. (D) (ii) Histogram showing the NND of all 55 tetramers identified in 5 similarly treated cells.    105 4.4 Discussion The current view of RYR2 disposition within the cardiac dyad is derived largely from scanning EM of skeletal muscle RYR1, coupled with two dimensional images of cardiac muscle dyads (Ferguson et al., 1984; Franzini-Armstrong et al., 1999). My results, which utilize the three dimensional capability of electron tomography, have provided views of the dyad and its associated RYR2 that could not have been previously obtained. I have shown that under physiological conditions the distribution of RYR2 within a dyad is neither homogenous nor well structured (Fig. 4.1) and that when the environment is altered the arrangement of the tetramers changes quickly (in minutes). The pseudo-crystalline array of RyR1 observed in lipid bilayers subjected to various concentrations of Mg2+ implies that the nearest neighbour distances (NND) between the tetramer centers within a dyad should (assuming the 29 nm square tetramer that I used in our calculations) be either 30.3 nm (side-by-side array) or 32.4 nm (checkerboard array) (Yin et al., 2005a). The latter value imposes an overlap of 14.5 nm between the sides of the RyR1, allowing interaction between adjacent tetramers’ clamp domains, should it occur5. The NND of the combined rat and human data (Fig. 4.3 and Fig 4.4 A-C) shows that even under normal physiological conditions the arrangement of the tetramers cannot be characterized using a simple two-state model. In addition, the distribution of the NND of those tetramers that could form a checkerboard arrangement had a roughly bell shape with a peak at 38 nm, a value far greater than that of the RYR1 model. Further, such a broad distribution implies that under normal physiological conditions the tetramers are not in any fixed position and although they often abut, their degree of overlap is highly variable, suggesting that the positioning of the tetramers is dynamic and that fixing the cell is providing a snapshot of this process. This contention is supported by my   106 observations of the effects of phosphorylation and varying the free Mg concentration, all of which led to a dramatic change in the positioning and the NND distributions compared with the controls (cells fixed in situ; cells isolated and fixed; cells isolated, permeabilized with the free Mg2+ set to 1 mM and the free Ca2+ to 100 nmol/l and then fixed). Notably, saponizing the sarcolemma had no effect on the results (Fig. 4.4 C). These findings are controversial and I therefore analyzed our methodology to determine whether the large value for the NND as well as the broad distribution could be artifacts arising from the identification, placement and measurement process.  Identifying the tetramers and differentiating them from membrane and other structures, although laborious, is straightforward when using the three orthogonal images as a guide, provides a general location for the position of a tetramer, and has a low error rate. The final positioning of the tetramer, by fitting the 29 nm square box on the en-face (XZ) view of the dyad, has an inherent error of about 1 nm due to the thickness of the line drawn on the screen. The positioning and orientation was often helped by the tetramers having a clear edge or corner and being adjacent to other tetramers. The latter situation was particularly useful since a single well-defined tetramer was used to seed the position of the adjacent tetramers given that they cannot overlap. Although our judgment of best fit was done by eye, it was constrained by these conditions. In addition, I have individually reviewed the positioning of the tetramers and found that differences rarely exceeded 3 nm. Even when the position was in dispute the large number of tetramers examined (over 600) meant that such errors would have little or no effect on the NND distributions. Heavy metal staining, a requirement for generating the electron tomographic image, sometimes gave the tetramer unclear boundaries and if there were no adjacent tetramers I positioned the tetramer in the center of the blob and guessed as to the orientation. Such situations   107 were also rare and had little influence on the final result given the large number of tetramers that I identified. The determination of the exact orientation of the receptor is the weakest part of this process, but because I used the center-to-center distances for calculating the NND, any uncertainty in the orientation has no effect on the values in the histograms or on the conclusions I reached.   I also analyzed the effect that curvature or the undulating nature of the dyad would have on these results. Our technique maps the tetramer positions onto a horizontal plane above the dyad, which is where I calculate NND, so the effect of a curvature would be to decrease the distances calculated, leading to an underestimate. However, the effect is minimized because tetramer clusters tend not to straddle a curve (see Fig. 4.13) and even if they do the error is a few percent at most, since the change in depth over the length of a dyad (250 nm or more) was never more than 25 nm, less than the width of a single tetramer.   Another possibility is that I have a scaling error and that the real size of boxes fitted to the tetramers is smaller than 29 nm. There are a number of arguments against this hypothesis: First the boxes fit the tetramers well and in many cases tetramers abut (see Fig 1, for example); any larger and they would overlap which is clearly impossible.  Second, the error would have to be consistent in each of the 52 tomograms collected as well as quite large (~ 20%), Last, the NND for the side-by-side tetramers is close to that predicted by Yin et al.11 suggesting that our scaling is reasonably accurate. There was a dramatic widening of the SR lumen in low Mg2+ (Fig. 4.8 D). This was not due to low osmotic pressure and simple cellular swelling as the mitochondrial cristae were clearly visible and well preserved. Since the peak NND (39 nm) was no different from that seen for tetramers in a checkerboard configuration in both the control (38 nm) and phosphorylated   108 cells (38 nm), the phenomenon cannot be due to an overall expansion of the SR as it would be expected to shift the peak NND to larger values. I hypothesize that structural elements holding the junctional SR membranes together require bound Mg2+, and in its absence the membranes separate and the lumen of the junctional SR widens. The changes that I observed in the distribution and its associated NND when phosphorylating the receptors or changing the free Mg2+ concentration also support our contention that the results in Fig 4.3 are not an artifact. Both phosphorylation (Fig 4.13 Dii) and low Mg2+ (Fig 4.7 D) reoriented the tetramers into a largely checkerboard formation with a peak NND of 39 nm, close to the 38 nm found in our controls, while high Mg2+ packed the tetramers into a mostly side-by-side formation with a peak of 30 nm (Fig 4.11 Cii), close to the Yin model for side-to-side tetramers (Yin et al., 2005a). The distribution of the tetramers under control conditions would seem to be a mix of these two extremes that could be explained by an intermediate Mg2+ concentration (1 mM) and basal phosphorylation of RYR2 (Huke and Bers, 2008).  All of these considerations led me to conclude that the large values and broad distribution of the NND of tetramers in a checkerboard arrangement, and the dramatic alterations in the distribution following changes in the environment, were real and evidence for the tetramers being both mobile and able to respond to changes in physiological and pathological stimuli by repositioning within the dyadic cleft. Importantly, tomograms of human myocytes produced qualitative and quantitative results that were indistinguishable from those obtained from the rat under control conditions (Fig. 4.5 and Fig. 4.6). It is therefore likely that human RYR2 are equally mobile and would respond to local changes in phosphorylation and Mg2+ concentration in a similar manner.   109 Comparing the tetramer organization with the spark frequencies, it is notable that stimuli which organized the tetramers into a largely checkerboard arrangement (low Mg2+ and phosphorylation) were associated with marked increases in the spark frequency compared to the control where the arrangement was mixed (Figs 4.1 Bi and Fig. 4.3). In contrast, a high Mg2+ concentration organized the tetramers into a side-by-side configuration (Fig 4.11 and Fig. 4.9) and was associated with a significant decrease in the spark frequency. The relationship is non-linear because phosphorylation or low Mg2+ doubles the proportion of tetramers in a checkerboard configuration compared to control, whereas the spark frequency increases about 6-fold. These results suggest that a correlation exists between the tetramers’ arrangement and their open probability, which would fit well with our observation in control cells of the tetramers being in a mixed configuration and giving rise to an intermediate spark frequency.  Although the 0.1 and 4 mM concentrations of Mg2+ are unlikely to occur in vivo, I have shown that the tetramers move in response to changing the Mg2+ concentration as well as to changes in the phosphorylation levels. A question that arises is why both phosphorylation and a low Mg2+ produce the same tetramer arrangement and are associated with an increase in the spark frequency. Li et al. observed that RYR2 phosphorylation produced a rightward shift in the Mg2+ concentration dependency of RYR2 inhibition, which they interpreted as a decrease in the tetramers’ affinity for Mg2+ (Li et al., 2013). If true, phosphorylation might reduce the amount of bound Mg2+, shifting the tetramers into a checkerboard arrangement. There is no obvious reason why the tetramers would shift their relative positions as they alter their open probability, but a possible explanation is the experimental observations of, and the theoretical models proposing, inter-protein allosteric interaction. Positive allosteric interaction has been reported between adjacent RYR2 (Marx et al., 2001a) and RYR1 (Porta et   110 al., 2012a) tetramers in vitro, and while the former used no Mg2+ in their solutions, the latter observed an inverse relationship between the free Mg2+ concentration and the degree of allosteric interaction. RYR2 phosphorylation has also been reported to increase the synchrony of Ca2+ release, among other actions (Ogrodnik and Niggli, 2010). These results, coupled to our own, imply that the checkerboard configuration is associated with more tightly co-ordinated channel openings and possibly with positive allosteric interaction. However, if there is positive allosteric interaction it is unlikely to be explained by a simple model involving a fixed position on the tetramer’s clamp domain since the degree of overlap between adjacent tetramers is highly variable. The correlation between low spark frequency and the side-by-side configuration may represent evidence for the theoretically proposed negative allosteric interaction (Stern et al., 1999), but remains speculative in the absence of any other confirming data. In conclusion, our results provide a new framework for investigating, understanding and modeling the function of the dyad.   111 5. General discussion 5.1 Overview  Our view of myocyte architecture has radically altered in the past few years, and it continues to evolve. In this thesis, I present a new view of dyad distribution and of the placement of ryanodine receptors within them. I have identified RyR2 orientation within couplons in normal physiological conditions and examined the effects of inhibitors and activators on CICR in cardiac myocytes. The strengths and advances of the work presented in this thesis have been: 1) to demonstrate that RYRs on axial junctions likely play a role in EC coupling; 2) to demonstrate that RyR2s do not form an ordered array, rather they are scattered throughout it; 3) to demonstrate that changes in the local environment have an effect on the RyR2 tetramers’ position within the couplon; 4) to reveal that individual tetramers can both rotate and translate independently of each other; and 5) to demonstrate that the distribution of RyR2 in normal human ventricular cells is comparable to what I have observed in rat heart tissue.   The function is a direct consequence of the structure. This work provides a foundation and a model for the future efforts to further dissect the ECC mechanisms in both normal and diseased heart. This is important because RyR2 mutations instigate cardiac disease in humans such as CPVT. Also, hypertrophy and heart failure both involve alterations in CICR. In this discussion, I will tie together the findings of chapter 2,3 and 4 that form the body of this thesis, while discussing the overall significance of the work and the directions that have been taken so far and will be taken in future to build upon this work.    112 5.2 Merging the finding of this thesis work and its significance to the field  5.2.1 Cellular architecture and calcium release units Ca2+ was established as the main intracellular messenger for cell contraction by the initial study, completed in frog hearts (Ringer, 1883). In a basic functioning model, the process from depolarization of the plasma membrane to contraction is the result of the spatial and temporal coordination of the molecules that release Ca2+ into and remove it from the myoplasm. My work has provided a role in renewing our understanding of the localization of the channels involved in ECC and their molecular components.  The link between electrical excitation of the membrane to contraction in muscles was established long before its molecular components, the ion channels and transporters on the sarcolemma and SR, were identified (Dulhunty, 2006). Later, electron microscopy provided valuable insights into muscle structure, first in skeletal and then in cardiac muscle. The fundamental research in Moore’s laboratory has been to map the molecular architecture of cardiac myocytes, with an emphasis on the molecules that are involved in EC coupling. Using established diffraction limited fluorescence imaging protocols and imaging analysis techniques, I showed that a substantial portion of the RyR2s, 40%, did not colocalize with Cav1.2 but did colocalize with Cav3 (Scriven et al., 2005). I hypothesized that these subpopulations of RyR2s, which I called extradyadic RyR were in a separate domain whose function was controlled by the signaling molecules bound to Cav-3, such as PKA, eNOS or β-AR. I also hypothesized that these RyR2s were located on cSR in close proximity to Cav-3 and that this association formed a unique signaling pathway for modulating Ca2+ release from the SR. However, after examination of more than 200 micrographs of cardiac myocytes I was not able to find any cSR in ventricular cells. I performed a parallel examination of atrial myocytes that   113 confirmed the extensive distribution of cSR in that tissue demonstrating that I could see cSR in ventricular if it was there. I was not able to find any caveolea on the TATS system (2D EM), which has been confirmed by other group who used 3D EM (Wong et al., 2013). Given that others have identified caveolae on the TATS, further investigation under normal and pathological conditions may be warranted (Levin and Page, 1980). My observation that there is no cSR and no caveolea in the t-tubules led to the hypothesis that RyR2 without adjacent Cav1.2 are in dyads and that Cav-3 is in the t-tubule membrane facing the RyR2 in jSR (Figure 5.1). Because those RyR2s might be located in a dyad, the “extra dyadic RyR” is not a proper term to use. I suggest that an appropriate term for the RyR2’s adjacent to Cav3, but not to Cav1.2, be “extra couplonic RyR”.  A recent investigation using both super-resolution immunofluorescence and 3D electron tomography has provided some evidence for this, finding that about 5% of RyR2s colocalize with Cav3 on the surface, and that this doubles in the cell interior (Wong et al., 2013). The structural significance of this association remains to be determined. Future studies will have to use super-resolution immunofluorescence whose resolution is ~30 nm (Betzig et al., 2006), or double-label immunoelectron microscopy. However, further examination of the structure and function of the “extra couplonic RyRs” was postponed because of our interesting finding concerning the unexpected distribution of RyR2s within a couplon (Chapter 3).     114  Figure 5.1 Summary schematic of RyR2 distribution within dyad and extra couplonic RyR2.   As stated above, RyR2s are almost exclusively associated with the sarcolemma and form dyads. My early work (Chapter 2) showed that many of the axial tubules have junctions and that these junctions are quite extensive, sometimes running all the way from one Z-line to the next. I found the axial junctions are structurally similar in a number of ways to those on the transverse tubules. First, the axial junctions have both RyR and CSQ, the distance between the axial tubule and the jSR membrane is only 10-15 nm, and the axial tubule membrane anchors Cav1.2 opposite RyR in the jSR, identical to the structure of the junctions found in the t-tubules. Second, the structural features of the axial junctions in atria are identical to those of the ventricle, and these atrial junctions are known to be active participants in ECC (Kirk et al., 2003). Third, the large and relatively frequent assemblies of RyR in the axial junctions are sufficient to explain some aspects of Ca2+ dynamics. Up to 20% of a rat ventricular myocyte’s spontaneous Ca2+ sparks originate from areas of the sarcomere that are too far from the Z-line to be attributed to t-tubule junctions (Lukyanenko et al., 2007; Shacklock et al., 1995). In addition, the immuno-staining of A"B"RYR2"tetramer"t"t"Cav-3  RyR jSR T-tubule Sarcolemma)Cav1.2 Β2 AR Axial-tubule   115 RyR and Cav1.2 (Chapter 2) showed levels of colocalization that were higher in the axial than the transverse junctions, suggesting the active involvement of axial junctions in ECC.  In addition, it has been difficult to account for the comparable horizontal and longitudinal conduction velocities of Ca2+ waves if the RyR are only on or near the Z-line (Subramanian et al., 2001). Again, my observations suggest that axial junctions might solve that problem. It is therefore reasonable to expect that axial junctions participate in normal ECC, as well as in the production of sparks and the spread of Ca2+ waves. However, the advantage of RyR2s distributed in axial junctions is not clear. Examination of rabbit neonatal cardiac myocytes showed a subpopulation of RyR2s distributed on the M band region, most likely adjacent to axial tubules, that gradually decreases in adult cells (Dan et al., 2007). The RyR2 can be repositioned during cardiac remodeling as for example in heart failure. Recent work has found a pathway for RyR2 turnover through chaperone-mediated autophagy that may be active during oxidative stress (Pedrozo et al., 2013). All together I speculate that RyR2 tetramers might assemble or turnover within axial tubules on the SR.   Studies based on a wide variety of animal models (mice, rats, dogs and pigs) have shown that heart failure (HF) is characterized by the disappearance of transverse tubules and their replacement by many in the axial direction. Studies on human tissue examining fixed or frozen samples, although uncommon, support the presence of similar changes in human heart failure (Zhang et al., 2013). The functional consequence of these structural changes is not clear, although it has been suggested by Sipido’s group that it can be linked to a loss of synchrony, as well as other functional changes in the time course of NCX-dependent Ca2+ removal (Biesmans et al., 2011). Overall, it is unclear whether or not the appearance of the axial tubules in HF is associated with the formation of axial junctions. Further, if these junctions are present, it is not   116 known what advantage or disadvantage they might provide to a failing heart. Both of these topics could be subjects for future investigation.    My first single tilt electron tomography examination of RyR2 channels was done on an axial junction (Chapter 2). The purpose of this study was largely to investigate the 3D structure of couplons and the associated t-tubules. I expected to see the checkerboard arrays of RyR2 in tissue fixed in situ.  The 3D model of the junction showed an unexpected disordered array of the tetramers. Since I only had one data set, we decided to publish the 3D structure showing only one layer of the RyR2 channels and concentrate on t-tubule geometry (Chapter 2). This finding was the basis of my further investigations. It is important to note that the conventional fixation method, using glutaraldehyde, was the only tissue fixation method that allowed me to visualize the cardiac cellular structure. The conventional fixation method in EM using glutaraldehyde has been thought to fail to preserve structural information about dynamic cellular components due to the slow processes of fixation. Even using a lab microwave for speeding up the processes, the fixation and dehydration takes minutes. The cryo-fixation technique overcomes this limitation as it fixes the cells very quickly, in 0.1ms (Sitte, 1996). I invested a fair amount of time examining different cryo-fixation methods (Immersion freezing, high pressure freezing etc.) with very little success. Whether the chemical fixation would affect the RyRs molecular structure and distribution is unknown. A recent study using single particle cryo-electron microscopy of RyR1 ultrastructure indicated that glutaraldehyde has no effect on the channel’s overall structure (Strauss and Wagenknecht, 2013). Although comparable structural studies of control RyR1 and glutaraldehyde-exposed RyR1 showed some conformational changes mostly on the clamp region where subunit 6 is located causing the channels to adopt an open-like conformation (Strauss and   117 Wagenknecht, 2013). All of the cells I examined are equally treated with gluteraldehyde, so it is unlikely that effects of the fixative would explain the large differences in RyR2 distribution that I observed under the various conditions. However, the checkerboard configuration is associated with a broad range of tetramer overlaps, and if RyR2 responds to gluteraldehyde, as does RyR1, then gluteraldehyde might open the clamp domain and allow the receptor to move prior to fixation. Such an effect could produce the broad range of overlaps that I see for the checkerboard configuration.   I have also contributed to work analyzing the distribution of RyR2 on a larger scale than can be seen using electron microscopy (Scriven et al., 2010). This work was done using diffraction-limited immunofluorescence. The lab initially showed that RyR2 couplons were about 1 µm apart in rat ventricular myocytes, which is roughly the width of a myofibril (Scriven et al., 2000b). Later analyses using more sophisticated analytical tools demonstrated that the NND of RyR2 couplons was 552 nm (Scriven et al., 2010). Other measurements gave comparable results, as they estimated the NND of 1.05 µm for RyR2 clusters on t-tubules compared to 0.83 µm on axial tubules in isolated ventricular myocytes (Izu et al., 2006), which was later confirmed by another group (Soeller et al., 2007).     5.2.2 Dyadic clefts are not filled with RyR2 It had been proposed that a dyad in a ventricular myocyte could be divided into a number of separate, smaller, units: “…the apparently single, large units in cardiac muscle may in reality be an aggregate of smaller, closely spaced units.” (Franzini-Armstrong et al., 1998). This idea has been confirmed in studies using electron tomography (Hayashi et al., 2009) and super-resolution immunofluorescence microscopy (Baddeley et al., 2009), both of which clearly   118 demonstrated that the dyad was not filled with RyR2, but was indeed ‘an aggregate of smaller, closely spaced units’. My work (Chapter 3) has also confirmed this view, although limitations of the 200KV Tecnai G2 TEM made measurement of the distance between dyads impossible since the thickest EM block that can be examined is between 200 nm and 300 nm, which can be smaller than one dyad. The concept of partially filled dyads is also supported by biochemical estimations of the RyR2 density in SRs. High affinity ryanodine binding study, has estimated eight RyR2s in a middle sized couplon that has an area that could hold ~43 RyRs (Hayashi et al., 2009). Figure 5.2 illustrates some examples of this sparse packing in three regions of the TATS system (i.e. axial, transverse and surface). I made 3D models of the dual tilt tomographic images of junctions using one of the initial modeling programs for tomography, known as IMOD. In 2009 the available free version of IMOD did not allow me to examine the en face view of the dual tilt and aligned tomograms, so making a model of the tilt series was the only available way to look at the distribution of the RyR2 channels on the surface of the jSR (Figure 5.2). Later in 2010, the Bio Imaging Facility of UBC purchased AMIRA, a modeling program with an advanced visualization option that enable me to view the surface structure of jSR and to identify the position of each tetramer in all three dimensions.      119  Figure 5.2 3D Electron tomography reveals the nano-scale 3-D structure of dyads in rat ventricular myocytes. High-resolution EM images of dyads in A) surface B) axial and C) transverse view; RyRs traced in pink, sarcolemma or TATS in green and jSR in blue. (ii) Surface mesh model of the junction from i. (iii, iv) Mapping RyRs in the junction shows that junctions are not filled with RyR2.The placement and orientation of individual tetramers could not be determined.  Scale bars, 50 nm.    5.2.3 RyR2 do not occur in regular arrays As described previously, couplons contain a cluster of Cav1.2 on the sarcolemma that is functionally coupled to RyR2 clusters on the jSR. Together they form functional Ca2+ release units.  A B C i ii iii iv i ii iii iv i ii iii iv   120 In chapter 3, based on an en face view of a dyad from a dual tilt tomogram, I showed that RyR2 distribution in the cleft might be a mixture of checkerboard and side-by-side arrangements. That conclusion was based on the inability to fit a regular checkerboard distribution to the data, it could only be fit with a mixture of side-by-side and checkerboard. I also observed some isolated channels that were physically uncoupled from other RyR2. The existence of these isolated channels, so-called ‘rogue’ RyR2s, had been suggested (Sobie et al., 2006) not long before my observations. It was hypothesized that ‘rogue’ RyR2s functioned differently and might act as a trigger for Ca2+ sparks (Sobie et al., 2006). While the limitation of our analytical techniques at that time prevented firm conclusions, I speculated that the two observed configurations might have differing allosteric interactions that could affect RyR2’s function, suggesting that there could be at least two functional groups of RyR2s within a couplon.  The new observations of dyad and RyR2 distributions from my work, and other groups, supported a model of the dyad in which the RyR2 are distributed in a number of small groupings including ‘rogue’ RyRs, with the tetramers having multiple arrangements; this contrasted with the earlier view of RyR2 distribution (Figure 5.3)   121  Figure 5.3 New and old view of dyad. Contrast between the previously held (A) and newly developed (B) views of RYR2 distributions within the dyadic cleft. jSR is shown in outline (light patches) with their complement of RYR2. t, lumen of t-tubules.   5.2.4 Non-uniform arrangements  In Chapter 4 of this thesis, I extended my earlier work using high-resolution dual tilt electron tomography, measuring the nearest neighbor distances (NND) within RyR2s and confirming that under physiological conditions (100 nM, Ca2+; 1 mM Mg2+) the distribution of RyR2 within a dyad is a mixture of largely checkerboard (peak NND ~39 nm) and side-by-side packing (peak NND of ~30 nm). I also found that when the environment is altered, the arrangement of the tetramers changes quickly: 1) high cytosolic Mg2+ changes the channel A"B"RYR2"tetramer"t"t"  122 distribution to a mostly packed side-by-side arrangement in vivo and 2) phosphorylation and low cytosolic Mg2+ changes the distribution to a largely checkerboard arrangement. Cytoplasmic magnesium The effect of cytosolic Mg2+ on channel distribution is important because Mg2+ is a known physiological regulator of RyR2. Cytoplasmic Mg2+ inhibition of RyR2 is shown to be due to Ca2+ competing with Mg2+ at the same binding sites (Laver and Honen, 2008). Lower Mg2+ increases, and elevating decreases, Ca2+ spark activity in saponin-permeabilized myocytes (Gusev and Niggli, 2008). As stated before, Mg2+ also decreases Ca2+ release from SR vesicles in response to 10 µM Ca2+ (Meissner and Henderson, 1987) and in situ evidence indicates that Mg2+ plays a significant role in determining RyR activity in vivo (Zahradnikova et al., 2010).   Given my results, I therefore speculated that in the normal physiological concentration of Mg2+, it was unlikely that all the channels had the same affinity or sensitivity to Ca2+. Channels in checkerboard arrangement might be more sensitive to Ca2+ than the group inhibited by Mg2+ that form the side-by-side arrangement. The correlation between spark frequency and RyR2 arrangement that I have shown in chapter 4 indicates that reorganization of the RyR2s may be another mechanism to regulate Ca2+ transport in cardiac myocytes. Together with my previous work, I have revealed that the RyR2 tetramers are not static, their position and packing changes in response to changes in the Mg2+ concentration. RyR1 crystallized in lipid bilayers in vitro and subjected to various concentrations of Mg2+ suggests that the nearest neighbor distances between the tetramer centers within a dyad should be either 30.3 nm (side-by-side configuration) or 32.4 nm (checkerboard configuration) with a 14nm overlap on clamp regions (Yin et al., 2005b). However, my observations within   123 ventricular myocytes indicating a wide-ranging distribution imply that under normal physiological conditions the tetramers are not in any fixed position and although they are often adjacent to each other, their degree of overlap is highly variable.  Several questions have still not been resolved. For instance, Mg2+ alters the orientation of the RyR2 tetramers, but what about other RyR2 inhibitors such as CaM or ryanodine? Do these inhibitors all function, at least in part, through a re-arrangement of the array to all side-by-side, or is it possible to inhibit RyR without changing the individual orientation? We expect with the use of electron tomography and the analysis program presented in this thesis along with the proper treatments, these fundamental questions can be answered in the future. The effect of phosphorylation on channel distribution It is well established that the RyR2 can be phosphorylated at a number of sites by different kinases like PKA and CaMKII, and that phosphorylation increases the open probability and plays an important role in arrhythmogenesis (Lokuta et al., 1995; Marx et al., 2000), but beyond that there is significant controversy. By phosphorylation of RyR2, directly, CaMKII is able to raise the Ca2+ spark frequency. On the other hand, phosphorylation of RyR2 by PKA indirectly affects spark frequency by elevating SR Ca2+ content (Vinogradova et al., 2006). A recent study of the crystal structure of the phosphorylation domain of RyR1 showed that the phosphorylation sites are all located at the clamp region (Yuchi et al., 2012). As mentioned earlier, the clamp regions go through large motions upon channel opening, therefore phosphorylation of RyR may affect allosteric motions in the region that normally accompanies channel opening (Yuchi et al., 2012). In Chapter 4, I showed that phosphorylation (the receptors may be hyperphosphorylated) changes the distribution of RyR2 mostly to a checkerboard   124 formation. It is currently unknown whether PKA and CaMKII equally collaborate in this reorientation. The phosphorylation cocktail I used activates endogenous PKA, as well as directly adding cAMP, and inhibits dephosphorylation by PP1 and PP2. In addition to activating PKA, cAMP directly activates Epac (exchange protein directly activated by c-AMP), which in turn activates CaMKII (Pereira et al., 2007), so both kinases will be active. This was the necessary first step. Future studies will be needed to determine whether one, or both, kinases must be active to rearrange the tetramers. In my initial attempt to answer this question, I activated endogenous CaMKII by increasing Ca2+-CaM in the presence of the PKA inhibitor, H89 (Guo et al., 2006). The results of Western blot analyses (Figure 5.4) demonstrate that the receptor had been highly phosphorylated on S2814 and not phosphorylated in the control samples. Using electron tomography (n =3) I found that phosphorylation by CaMKII alone did not change the orientation and that the center-to-center NND between the tetramers is comparable to the control (Figure 5.5). It is difficult to ensure whether the absence of any structural change after CaMKII treatment is a real phenomena or it is a result of the complex interactions that occur between CaM and RyR2. Ca2+-CaM used in my treatment has a high affinity for RyR2 and also an inhibitory effect (Ikemoto et al., 1995). The lack of change in the channel distribution may be due to the conflicting effects of activation by CaMKII phosphorylation and inhibition by Ca2+-CaM. Earlier studies showed that the activation of endogenous CaMKII has an inhibitory effect on RyRs that were already dephosphorylated by PP1 (Hain et al., 1995). It is possible that channels with a basal PP1 dephosphorylation do not show any change in the orientation. By continuing studies on the effect of exogenous CaMKII and PKA on channel orientation and using proper inhibitors to eliminate other factors (i.e. SR Ca2+or de-phosphorylation), we will get a better understanding   125 of the mechanism that controls RyR2 orientation. We also do not yet know whether the reorientation I have observed requires the tetramers to be phosphorylated, or perhaps an element of the cytoskeleton that moves the tetramers, or both.         126  Figure 5.4 Phosphorylation of ser-2814 in response to increase of endogenous CaMKII in rat cardiac myocytes.  Freshly isolated rat cardiac myocytes were stimulated by CaMKII Cocktail (CaM; 1.4 µM, Ca2+; 500 nM) or K93 (CaMKII inhibitor); 1µM for 10 min. Method adapted from (Guo et al., 2006). The chart demonstrates the relative phosphorylation in intact, saponin permeablized myocytes, treated with CaMKII cocktail and CaMKII cocktail plus K93.   Control,	  Intact CaMKII	  cocktail CamKII+K93 RYR-­‐S2814 Saponin	   0	  0.5	  1	  1.5	  2	  2.5	  Control,Intact	   Saponin	   CamKII-­‐endo	   CamKII+K93	  Rela?ve	  RYR-­‐S2814	  	  	  phosphoryla┱n	  RyR2	  Phosphoryla?on	  (N=5)	  360-­‐ MW kDa-­‐ 268-­‐   127    Figure 5.5 RyR2 distribution in a permeabilized rat cardiomyocyte with activated endogenous CaMKII.  (A) Orthogonal views, (i) XY, (ii) YZ, (iii) XZ, 1 nm thick, through a dyad, with the intersecting planes positioned over a single RYR2. Double arrow, t-tubule; single arrow, sarcoplasmic reticulum. (iv) tetramers.  (B) Histogram showing the NND of tetramers. n=3     i ii iii  50nm  iv A 0	  2	  4	  6	  8	  10	  12	  14	   28	    30	    32	    34	    36	    38	    40	    42	    44	    46	    48	    50	  Checkerboard	   Side	  by	  side	   Isolated	  NND	  #	  Tetramers B   128 5.2.5 Channel movement In Chapter 4, I showed that individual tetramers tend to reorient themselves within the membrane in response to regulators. What is the mechanism underlying this reorientation? Is there another regulatory protein controlling these movements? The cytoskeletal proteins tropomyosin 5 (TM5) and tropomyosin 4 (TM4) are located in the dyadic cleft area of skeletal muscles and are required for normal ECC by forming a protein network that organizes the localization of junctional proteins (Vlahovich et al., 2009). Another crucial study is to test whether cytoskeletal proteins like TM4 or TM5 have a direct regulatory effect on RyR2 reorientation.   5.3 Significance  All together, the work of this thesis provides a new view of dyad distribution and of RyR2 distribution within the dyad. I showed that RyR2 clustering is not static, individual tetramers can be rearranged, and that this is correlated with a change in their open probability.  My findings provide potential new mechanisms by which the activity of RyR2 tetramers, the dyad and cardiac contractility may be regulated. In the future it may be possible to manipulate the structure to achieve a desired functional goal, but we are not yet close to that ideal. Before then, we must first know what that structure is, and understand how the structure and its variations give rise to function.     129 Bibliography  Acsai, K., Antoons, G., Livshitz, L., Rudy, Y., and Sipido, K.R. (2011). 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