Modulation of Ca2+ Signaling by

Trimeric Intracellular Cation Channels in the Heart

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of the Ohio State University

By

Xinyu Zhou

Ohio State University Biochemistry Program

The Ohio State University

2019

Dissertation Committee

Dr. Jianjie Ma, Advisor

Dr. Jill Rafael-Fortney

Dr. Hua Zhu

Dr. Jian-Qiu Wu

Copyrighted by

Xinyu Zhou

2019

Abstract

Trimeric intracellular cation channels, called TRIC-A and TRIC-B, are distributed to intracellular Ca2+ stores and mainly mediate the permeability of K+ ions in multiple cell types. Previously we showed that genetic ablation of TRIC-A leads to compromised K+- permeability and Ca2+ release in the muscle sarcoplasmic reticulum (SR), supporting the hypothesis that TRIC channels function as counter-ion channels during the acute phase of

Ca2+ release under physiological conditions. In cardiomyocytes, spontaneous Ca2+ waves, triggered by store overload-induced Ca2+ release (SOICR) mediated by the type 2 ryanodine receptor (RyR2), develop extra-systolic contractions often associated with arrhythmic events.

In this study, we found that the carboxyl-terminal tail domain of TRIC-A (CTT-A) interacts with the RyR2 channel to directly modulate SOICR activity. Biochemical studies demonstrate direct interaction between CTT-A and RyR2. In HEK293 cells with stable expression of RyR2, transient expression of TRIC-A, but not TRIC-B, leads to apparent suppression of spontaneous Ca2+ oscillations. Ca2+ measurements using the cytosolic indicator Fura-2 and the ER luminal store indicator D1ER suggest that TRIC-A enhances

Ca2+ leak across the ER membrane by directly targeting to RyR2 to modulate SOICR.

Moreover, synthetic CTT-A peptide facilitates RyR2 channel activity in the lipid bilayer

ii reconstitution system and induces intracellular Ca2+ release after micro-injection into isolated cardiomyocytes, whereas such effects were not observed with the CTT-B peptide.

Therefore, in addition to the ion-conducting function, TRIC-A seems to function as an accessory protein of RyR2 to modulate SR Ca2+ handling in cardiac muscle.

Although Tric-a-/- mice do not display significant abnormity in the heart under resting condition, isoproterenol induced stress condition would induce abnormal cardiac function and fibrosis. Moreover, Tric-a-/- heart is more susceptible to Transverse Aortic

Constriction (TAC) induced heart injury and develop cardiac fibrosis and hypertrophy.

These results suggested that the stress resistance capacity is significantly compromised in

Tric-a-/- heart. Using isolated cardiomyocytes, we indentified altered SR Ca2+ uptake as well as altered mitochondria Ca2+ uptake in the Tric-a-/- cardiomyocytes indicating that

TRIC-A played a role in SR-mitochondria Ca2+ cross-talk.

In summary, our results demonstrate the crucial role of TRIC-A in regulating both

SR and mitochondria Ca2+ homeostasis. TRIC-A could directly interact with RyR2 and regulate its activity to maintain normal SR Ca2+ homeostasis. Cardiomyocytes lacking

TRIC-A would not only cause impaired SR Ca2+ signaling, but also affect mitochondria

Ca2+ homeostasis through SR-mitochondria cross-talk which leads to compromised mitochondria stability under stress condition and ultimately results in cardiac fibrosis and hypertrophy.

iii

Acknowledgments

I would like to present my deepest gratitude to Professor Jianjie Ma, for his guidance and encouragement throughout my PhD period. I would like to thank him for all of his hard work in mentoring me and helping me with this thesis. I would also like to convey my thanks to the members of Dr. Ma’s group for their help in this project and also for their friendship. Special thanks to Dr. Ki Ho Park, Dr. Pei-hui Lin, the experimental data of the TRIC-A project and Dr. Zehua Bian, Dr. Qiang Wang, and Dr. Hua Zhu for the

MG53 project.

I would like to thank my committee members, Dr. Jill Rafael-Fortney, Dr. Hua Zhu, and Dr. Jian-Qiu Wu, who provided valuable guidance and feedback for this thesis. Thanks also to the Ohio State Biochemistry Program for the kind guidance and support. Thanks to

Dr. Brian Foster for his review and suggestions for this thesis.

Finally, I would like to thank my wonderful wife, Jing Tan, for the constant love, encouragement, and support during my Ph.D. study. Thanks also to all of my family, especially to my parents, Zhijian Zhou and Fengyun Zheng, for their great support and unconditional love and trust.

iv

Vita

2009...... B.S. Northwest University, Xi’an, China

2011-2012 ...... Graduate Fellowship, UMDNJ/Rutgers, New

Brunswick, NJ

2012-Present ...... Graduate Research Associate, Department of

Surgery, the Ohio State University,

Columbus, OH

Publications

1. Adesanya T, Russell, M, Park KH, Zhou X, Sermersheim M, Gumpper K, Koenig

S, Tan T, Whitson B, Janssen P, Lincoln J, Zhu H, Ma J. MG53 protein protects

aortic valve interstitial cells from membrane injury and fibrocalcific remodeling. J

Am Heart Assoc. 2019.

2. Wang Z, Chen K, Han Y, Zhu H, Zhou X, Tan T, Zeng J, Zhang J, Liu Y and Li

Y. Irisin Protects Heart Against Ischemia-Reperfusion Injury Through a SOD2-

Dependent Mitochondria Mechanism. Journal of cardiovascular pharmacology.

2018.

vi

3. Fan Z, Xu Z, Niu H, Gao N, Guan Y, Li C, Dang Y, Cui X, Liu XL, Duan Y, Li H,

Zhou X, Lin PH, Ma J and Guan J. An Injectable Oxygen Release System to

Augment Cell Survival and Promote Cardiac Repair Following Myocardial

Infarction. Scientific reports. 2018;8:1371.

4. Elbaz M, Ahirwar D, Xiaoli Z, Zhou X, Lustberg M, Nasser MW, Shilo K and

Ganju RK. TRPV2 is a novel biomarker and therapeutic target in triple negative

breast cancer. Oncotarget. 2018;9:33459.

5. Fan Z, Fu M, Xu Z, Zhang B, Li Z, Li H, Zhou X, Liu X, Duan Y, Lin PH, Duann

P, Xie X, Ma J, Liu Z and Guan J. Sustained Release of a Peptide-Based Matrix

Metalloproteinase-2 Inhibitor to Attenuate Adverse Cardiac Remodeling and

Improve Cardiac Function Following Myocardial Infarction. Biomacromolecules.

2017;18:2820-2829.

6. Yao Y, Zhang B, Zhu H, Li H, Han Y, Chen K, Wang Z, Zeng J, Liu Y, Zhou X

and Wang X. MG53 permeates through blood-brain barrier to protect ischemic

brain injury. Oncotarget. 2016;7:22474.

7. Li H, Duann P, Lin PH, Zhao L, Fan Z, Tan T, Zhou X, Sun M, Fu M, Orange M,

Sermersheim M, Ma H, He D, Steinberg S, Higgins R, Zhu H, John E, Zeng C,

Guan J, e Ma J. Modulation of wound healing and scar formation by MG53-

mediated cell membrane repair. Journal of Biological Chemistry, 2015. 290, 24592-

24603.

vii

8. Ma H, Liu J, Bian Z, Cui Y, Zhou X, Zhou X, Zhang B, Adesanya TM, Yi F, Park

KH, Tan T, Chen Z and Zhu H. Effect of metabolic syndrome on mitsugumin 53

expression and function. PLoS One. 2015;10:e0124128.

9. Liu XY*, Zhou XY*, Hou JC, Zhu H, Wang Z, Liu JX and Zheng YQ. Ginsenoside

Rd promotes neurogenesis in rat brain after transient focal cerebral ischemia via

activation of PI3K/Akt pathway. Acta Pharmacol Sin. 2015;36:421-8.

10. Duann P, Li H, Lin P, Tan T, Wang Z, Chen K, Zhou X, Gumpper K, Zhu H,

Ludwig T, Mohler PJ, Rovin B, Abraham WT, Zeng C and Ma J. MG53-mediated

cell membrane repair protects against acute kidney injury. Sci Transl Med.

2015;7:279ra36.

11. Zhou X, Lin P, Yamazaki D, Park KH, Komazaki S, Chen SR, Takeshima H and

Ma J. Trimeric intracellular cation channels and sarcoplasmic/endoplasmic

reticulum calcium homeostasis. Circ Res. 2014;114:706-16.C

12. Park KH, Weisleder N, Zhou J, Gumpper K, Zhou X, Duann P, Ma J and Lin PH.

Assessment of calcium sparks in intact skeletal muscle fibers. J Vis Exp.

2014:e50898.

13. Jia Y, Chen K, Lin P, Lieber G, Nishi M, Yan R, Wang Z, Yao Y, Li Y, Whitson

BA, Duann P, Li H, Zhou X, Zhu H, Takeshima H, Hunter JC, McLeod RL,

Weisleder N, Zeng C and Ma J. Treatment of acute lung injury by targeting MG53-

mediated cell membrane repair. Nat Commun. 2014;5:4387.

viii

Fields of Study

Major Field: Ohio State Biochemistry Program

ix

Table of Contents

Abstract ...... ii

Acknowledgments ...... iv

Vita ...... vi

Table of Contents ...... x

List of Figures ...... xv

Chapter 1. Introduction ...... 1

Cardiac calcium signaling ...... 1

Calcium signaling ...... 1

Excitation-Contraction Coupling (ECC)...... 2

Store-overload induced Ca2+ release (SOICR) ...... 3

Regulation of excitation-contraction...... 4

Ryanodine Receptor (RyR) ...... 5

Molecular characterization ...... 5

Distribution of RyR2 in the cardiomyocyte ...... 7

Regulation of RyR2 ...... 8

Calcium, magnesium and ATP ...... 8

RyR2 regulatory proteins ...... 9

SR proteins: calsequestrin, triadin and junctin ...... 13

The need for countercurrents ...... 14

x

Trimeric Intracellular Cation Channels (TRIC) ...... 15

Identification of TRIC...... 15

Biochemical and biophysical properties of TRIC channels ...... 17

Role of TRIC-A in SR Ca2+ homeostasis in skeletal and smooth muscles ...... 18

Genetic variations of TRIC and their roles in human diseases ...... 21 Contribution of TRIC-A and TRIC-B to Ca2+ signaling in cardiac development and contraction...... 21

Chapter 2. TRIC-A Channel Maintains Store Calcium Handling by Interacting with Type 2 Ryanodine Receptor in Cardiac Muscle...... 30

Introduction ...... 30

Methods ...... 34

Animals ...... 34

Cardiomyocyte isolation and Ca2+ spark measurement ...... 34

Plasmid construction ...... 35

Single-cell Ca2 + imaging ...... 35

ER Ca2+ Measurement ...... 36

Immunoblotting...... 36

Immunofluorescence staining ...... 37

GST-CTT-A protein expression and beads binding assay ...... 37

Channel activity assay on planar bilayer system ...... 38

Microinjection of CTT-A and CTT-B peptide...... 39

Statistical analysis ...... 39

Results ...... 40

Abnormal Ca2+ signaling in cardiomyocytes derived from the Tric-a-/- mice ...... 40

Co-expression of TRIC-A and RyR2 in HEK293 cells modulates SOICR ...... 41

xi

TRIC-A interacts with RyR2 through its carboxyl-terminal tail (CTT) domain .. 43

TRIC-A carboxyl-terminal tail peptide increases RyR2 channel activity ...... 44

Discussion...... 47

Chapter 3. TRIC-A Regulates Cross-Talk of Ca2+ Between SR and Mitochondria in Cardiac Muscle...... 62

Introduction ...... 62

Methods ...... 65

Electrocardiography ...... 65

Transverse Aortic Constriction on Mouse ...... 65

Electron microscopy (EM) ...... 65

Cardiomyocyte isolation and Ca2+ spark measurement ...... 66

Immunoblotting...... 67

Histological analysis ...... 67

Results ...... 69

Tric-a-/- mice showed abnormal cardiac function and developed cardiac fibrosis under stress conditions...... 69 Additional ablation of TRIC-B in Tric-a-/- mice showed a much severer cardiac phenotype under normal and stressed conditions...... 70

SR Ca2+ signaling is altered in Tric-a-/- neonatal cardiomyocytes ...... 70

SR Ca2+ signaling is altered in Tric-a-/- adult cardiomyocytes...... 71

The heart of Tric-a-/- mice is more susceptible to TAC-induced stress...... 72 Mitochondria of Tric-a-/- cardiomyocytes showed more damage under the TAC induced stress ...... 73 Mitochondria Ca2+ signaling is altered in Tric-a-/- cardiomyocytes via cross-talk to altered SR Ca2+ signaling...... 74

xii

Mitochondria of Tric-a-/- cardiomyocytes are susceptible to acute oxidative stress while treatment of mitochondria Ca2+ uptake inhibitor RU360 would protect against damage...... 75

Discussion...... 77

Proposed model that TRIC-A regulates SR Ca2+ homeostasis and the Ca2+ signaling cross-talk between SR and mitochondria...... 77

Chapter 4. Sustained Elevation of MG53 in the Bloodstream Increases Tissue Regenerative Capacity without Compromising Metabolic Function ...... 89

Introduction ...... 89

Methods ...... 91

Experimental Animals ...... 91

Serum MG53 quantification ...... 92

Glucose Tolerance Test...... 93

Insulin Tolerance Test...... 94

Ear punch injury ...... 94

Treadmill running experiment ...... 95

Histopathology and Immunofluorescent Staining ...... 95

Immunoblotting...... 96

Cardiotoxin induced skeletal muscle injury ...... 97

High fat diet feeding ...... 97

Echocardiography ...... 97

Statistical Analysis ...... 98

Results ...... 98

Mice with sustained elevation of MG53 in the bloodstream lived a healthy lifespan ...... 98 tPA-MG53 mice show normal glucose handling and insulin signaling in skeletal muscle ...... 100 xiii

The tPA-MG53 mice show increased healing capacity following tissue injury . 102 Increased muscle performance and injury-regeneration with the tPA-MG53 mice ...... 103 Crossing of tPA-MG53 with db/db mice did not alter insulin signaling nor glucose handling...... 105

Discussion...... 107

Chapter 5. Conclusion and Perspective ...... 118

Summary of findings...... 118

Future work ...... 122

References ...... 130

xiv

List of Figures

Figure 1.1 EC-coupling ...... 24

Figure 1.2 CICR and SOICR...... 25

Figure 1.3 Structure of RyR2...... 26

Figure 1.4 Ca2+ /Mg2+ regulatory sites on the RyR2...... 27

Figure 1.5 Identification of TRIC ...... 28

Figure 1.6 Model for TRIC function in Ca2+ signaling ...... 29

Figure 2.1 Compromised Ca2+ spark signaling in cardiomyocytes derived from Tric-a-/- mice...... 52

Figure 2.2 Effect of TRIC-A and TRIC-B on RyR2-mediated SOICR in HEK293 cells. 53

Figure 2.3 Effect of TRIC-A on changes in ER Ca2+ load require interaction with RyR2.

...... 55

Figure 2.4 Co-localization of TRIC-A and RyR2 ...... 57

Figure 2.5 CTT-A interacts with RyR...... 58

Figure 2.6 Chimeras of TRIC-A and TRIC-B show different effects on SOICR in HEK293 cells...... 59

Figure 2.7 CTT-A directly modulate RyR2 channel function...... 60

Figure 3.1 Abnormal heart function in Tric-a-/- mice...... 80

Figure 3.2 Iso-induced bradycardia and sudden death in Tric-a-/- Tric-b+/- mice...... 81

Figure 3.3 Altered Ca2+ signaling in isolated Tric-a-/- neonatal cardiomyocytes...... 82

Figure 3.4 Altered Ca2+ signaling in isolated Tric-a-/- adult cardiomyocytes ...... 83 xv

Figure 3.5 TAC induced hypertrophy and fibrosis in Tric-a-/- heart...... 84

Figure 3.6 Increased mitochondria damage in Tric-a-/- heart after TAC...... 85

Figure 3.7 Altered SR and mitochondria Ca2+ signaling in Tric-a-/- cardiomyocytes after

TAC...... 86

Figure 3.8 Acute oxidative stress induces more mitochondria damage in Tric-a-/- cardiomyocytes...... 87

Figure 3.9 Model that TRIC-A regulates SR Ca2+ homeostasis and the Ca2+ signaling cross- talk between SR and mitochondria...... 88

Figure 4.1 Mouse model with sustained elevation of MG53 in the bloodstream...... 111

Figure 4.2 Assessment of insulin signaling and glucose handling in tPA-MG53 and WT mice...... 112

Figure 4.3 tPA-MG53 mice show increased healing capacity following ear-punch injury.

...... 114

Figure 4.4 Increased muscle performance of the tPA-MG53 mice under stress conditions.

...... 115

Figure 4.5 tPA-MG53 skeletal muscle showed enhanced regeneration capacity after cardiotoxin injury...... 116

Figure 4.6 Cross of tPA-MG53 mice with db/db mice did not alter the diabetic phenotype.

...... 117

Figure 5.1 Immunofluorescent staining showed TRIC-A is expressed on the nuclear envelope of cardiomyocytes...... 127

xvi

Figure 5.2 TAC significantly reduced invagination of nuclear envelope in Tric-a-/- cardiomyocytes...... 128

Figure 5.3 Immunofluorescent staining showed TRIC-A is highly expressed in mouse brain...... 129

xvii

Chapter 1. Introduction

Cardiac calcium signaling

Calcium signaling

Ca2+ ions are important second messengers in many biological signaling pathways, including muscle contraction, neurotransmission, cell differentiation, cell proliferation and apoptosis1-6. To regulate these many different pathways, the Ca2+ signaling systems show great dynamic range in terms of frequency, amplitude and spatial-temporal position, forming a variety of patterns from localized brief Ca2+ bursts to longer-lasting, global Ca2+ transients. On the other hand, sustained elevation of intracellular Ca2+ are known to be cytotoxic which would lead to mitochondria damage, dysregulation of gene expression, apoptosis and cell death, thus the physiological Ca2+ homeostasis and signaling must be tightly regulated. To assure that, there are many molecules and proteins including signaling ligand and receptors, ion channels, ion pumps, and regulatory proteins that are coupled together and are functioning in a coordinated manner7,8. Compromise of such Ca2+ homeostasis and signaling have been linked to different human diseases, including muscle dysfunction and heart failure1-5.

1

Excitation-Contraction Coupling (ECC)

In the heart, the electrical excitation signal needs to be transduced into mechanical contraction. Ca2+ act as a vital signaling messenger that couples those two processes together. This mechanism is well known as Excitation-Contraction Coupling (ECC, Figure

1.1)3. During the diastole phase (resting), intracellular Ca2+ is mainly stored inside of sarcoplasmic reticulum (SR) of the cardiomyocytes. During systole phase (activating) , the

ECC is initiated by the electrical excitation on the membrane of myocyte sarcolemma, in form of an action potential (AP), via the depolarization (from ~-80 mV to ~+40 mV) of cardiomyocyte plasma membrane3. The propagation of AP travel through plasma membrane into transverse tubules (T-Tubules), which are invaginations of cell membrane into the cells forming a complex membrane network for the ECC9. L-type Ca2+ channel

(LTCC) is the main type of the voltage-gated calcium channels located on the T-Tubule characterized by higher voltage activation10,11.

The invagination of the T-tubules brings the LTCCs in proximity to clusters of ryanodine receptors type 2, RyR2) on the junctional SR membrane12. As mentioned previously, AP passes through the sarcolemma membrane into the T-Tubules membrane and causes coordinated opening of LTCC leading to the influx of Ca2+ into the cardiomyocyte. This initial influx of Ca2+ is not substantially sufficient to trigger the contraction of cardiomyocyte, but it is enough to activate ryanodine receptor (RyR) channels located in the SR membrane that subsequently cause the efflux of Ca2+ from the

2

SR12,13. This elevated Ca2+ diffuses rapidly and further activates remaining RyR2 clusters through positive feedback amplification, leading to a global elevation of intracellular Ca2+, constituting a Ca2+ transient9,10,14-16. This process of initial small Ca2+ influx via the LTCCs on the t-tubule membrane triggering global intracellular Ca2+ release through RyR2 on SR membrane is known as Ca2+-induced Ca2+-release (CICR)8,14-16. The global Ca2+ elevation through LTCCs/RyR2 mediated CICR during systole leads to actin-myosin cross-bridge formation and ultimately muscle contraction17.

At the end of systole, depletion of SR Ca2+ content breaks the positive feedback of

CICR and causes the termination of Ca2+ release. During diastole, Ca2+ is depleted from cytosol opposite from systole that involve four types of process: SR Ca2+ uptake by Ca2+-

ATPase pump (SERCA)18, Plasma membrane Ca2+ extrusion by Na+-Ca2+ exchanger

(NCX) and Plasma membrane Ca2+-ATPase (PMCA) and mitochondrial Ca2+-uptake by mitochondrial calcium uniporter (MCU)14. Among them, the majority of cytosolic Ca2+ undergoes rapid reuptake into SR by SERCA in preparation of the next wave of CICR11.

Store-overload induced Ca2+ release (SOICR)

Ca2+ release from the SR in cardiomyocytes is normally controlled by voltage- dependent Ca2+ influx via the L-type Ca2+ channel through the CICR mechanism14. In addition to CICR, it has long been recognized that SR Ca2+ release can occur spontaneously under conditions of SR Ca2+ overload19-22. Considering its dependence on SR Ca2+ load and independence on membrane depolarization, this spontaneous SR Ca2+ release has been referred to as store-overload-induced Ca2+ release (SOICR)23,24,4 (Figure 1.2). When the

3

SR Ca2+ store was overloaded due to either external stress stimulus or internal disfunction of Ca2+ singling control, RyR2 could be activated by SR luminal Ca2+ rather than action potential triggered CICR. This SOICR events in the form of Ca2+ waves can enhance the activity of the Na+/Ca2+ exchanger, leading to delayed afterdepolarizations (DADs) and triggered activities3,25,26. These SOICR-evoked DADs and triggered activities are a major cause of ventricular tachyarrhythmias and sudden death in patients with catecholaminergic polymorphic ventricular tachycardia (CPVT) and heart failure3,25.

Regulation of excitation-contraction

During exercise or under external stress, both rate and strength of heart beat are greatly enhanced in order to accommodate the increased metabolic need of the whole body.

This change is mediated by β-adrenergic regulation. During β-adrenergic stimulation, catecholamines such as norepinephrine and epinephrine are release from sympathetic nerves and the adrenal medulla into the circulation system and bind to b-adrenergic receptors of cardiomyocytes. This will trigger downstream activation of protein kinase A

(PKA)-mediated phosphorylation of LTCC, PLN, RyR2, troponin I, and myosin binding protein C. PKA mediated phosphorylation thus systematically enhance the overall ECC signaling. For example, the phosphorylation of LTCC significantly increases Ca2+ entry during e-c coupling. The phosphorylation of RyR2 promotes the activation of the channel, which increase the SR Ca2+ release during each CICR. The phosphorylation of PLN diminishes its inhibitory effect on SERCA, leading to increased SR Ca2+ uptake rate and

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total SR Ca2+ content. The increased Ca2+ entry and SR Ca2+ load thus significantly enhance

Ca2+ transient amplitudes, leading to elevated contraction.

Ryanodine Receptor (RyR)

The Ryanodine Receptor (RyR) is one of the most important elements in ECC.

Ryanodine Receptors (RyRs) are Ca2+ sensitive high conductance Ca2+ release channels located on junctional SR and closely opposed to L-type Ca2+ channels on T-tubules in the muscle27. The name of RyR came from its ability to bind the ryanodine, a poisonous diterpenoid naturally in the roots and stems of South American plant Ryania speciose.

Ryanodine interacts with RyR with high affinity and specificity at open state of RyR. At low nanomolar concentrations of Ryanodine level, the RyR is locked in a permanent-open subconductance state, while at higher micromolar concentrations, activity of RyR channel is completely blocked. These features of Ryanodine make it a useful tool for RyR channel research28.

Molecular characterization

RyR2, known as the largest ion channel, is a homo-tetramer with each subunit at the molecular size of 560 kDa29. Three isoforms of RyR have been identified on mammalian cells: RyR1, RyR2 and RyR3. Around 70% of sequence identity is conserved in all three isoforms. RyR1, also known as skeletal isoform, is highly expressed in skeletal muscle30. It is also expressed at lower levels in vascular smooth muscle31, Purkinje cells,

5

cerebellum, stomach, spleen, kidney, testis, ovaries, adrenal glands32. RyR2, known as cardiac isoform, is the predominant expressed in cardiac muscle33. Moreover, RyR2 is also expressed in brain, in cerebellum and cerebral cortex at high levels 34, and in the stomach, lungs, kidney, adrenal glands, thymus and ovaries at low levels 32. RyR3 shows a wider distribution pattern in different tissues32,35.

One can imagine that the massive size and dynamic nature of RyR created significant difficulty in determining its molecular structure in the past. Bioinformatic analysis of RyR sequence revealed several domains that are similar with other proteins 29.

Only a few short domains were available for crystal structure, such as N-terminal fragments36-39, the phosphorylation sites of RyR40,41 and FKBP12 binding domain42.

Thanks to the recent “revolution in resolution” of single-particle cryo-EM, research from three independent group solved the detailed structural of RyR1 at the closed state with resolution43-45. These studies have provided information regarding the global architecture and conformational variations, as well as the detailed structure of functional domains such binding sites of regulatory molecules and accessary proteins for RyR channels46,47. The overall RyR structure is usually described as mushroom-like shape. (Figure 1.3) The big cytosolic “mushroom cap” portion consists of the N-terminal domain and around 80% of the entire protein, and the transmembrane and luminal portion consists of the C-terminal domain and around 20% of the entire protein42. The specific domain was termed differently from the three groups, but the general structure of channel was reported similarly43.

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Distribution of RyR2 in the cardiomyocyte

RyR2s are mainly found in the junctional SR, but they are also found in the corbular

SR (extra-junctional SR), serving as an additional amplification system to Ca2+ release from the dyads region48. However, early mathematical models of Ca2+ propagating simulation suggested that the calculated spatiotemporal Ca2+ distribution should lead to inefficient activation of muscle contraction49. This conflict, which has been called “the calcium paradox”, was solved by introduction of extra Ca2+ releasing sites across the longitude of the sarcomere49. Based on findings from super-resolution imaging50-52, EM tomography53,54 and 3D reconstruction studies48,55, researchers has suggested the existence of isolated RyR and non-uniformed small cluster of RyR that constitute the additional Ca2+ release sites. Experimental data also suggest one fifth of the total RyR2s are located outside the Z-line12,13,56, which is consistent with the previous mathematical estimation49.

It has been reported that RyR presents on perinuclear mitochondria56 and in SR proximal to the nuclear envelope57,58. Researchers have been proposed that these RyR2s could play certain roles in mitochondrial and nuclear signaling, as well as Ca2+-dependent gene regulation56,58,59. RyR2 distribution is different in neonatal cardiac myocytes and adult atrial because in these cells to lack of well-organized T-Tubule system, RyR2s are located at the periphery SR immediately under the sarcolemma. Thus compared to ventricular myocytes, these two cell types show some different patterns in Ca2+ signaling60.

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Regulation of RyR2

RyR2 exists in SR membrane in complex with variety of regulatory accessory proteins. Molecule ligand such as Ca2+, Mg2+ and ATP directly regulate RyR2 activity29,61.

Calcium, magnesium and ATP

Ca2+ is apparently the most important ion involved in the function of RyR2. Ca2+ can bind to the cytosolic and luminal sides of RyR2 to regulate the activation and inhibition of the channel activity. Compared to RyR1, RyR2 has a more potent response to Ca2+ induced activation yet a decreased sensitivity to millimolar Ca2+ inhibition. For RyR2, Ca2+ is the primary physiological trigger during the process of CICR is cytosolic. Under diastolic conditions (<100 nM), there is almost no spontaneous activation of RyR2. During systole, when cytosolic Ca2+ reaches to micromolar concentrations (1-10 µM), the RyR2 channel is activated. Further enrichment of cytosolic Ca2+ at millimolar concentrations (>10 mM) inactivate the channel62. Besides the cytosolic Ca2+ regulation mechanism of RyR2, several mechanisms of luminal Ca2+ regulation of RyR2 activity have been proposed. The first hypothesis is that RyR2 contains a luminal regulatory site that Ca2+ can directly bind to and regulate the activity of the channel63. The second hypothesis is that the regulatory mechanism is through the cross-talk of luminal and cytosolic Ca2+ binding site64. The third hypothesis is that the ‘luminal Ca2+ sensor’ of RyR2 is regulated through the interaction of calsequestrin/junctin/triadin complex with RyR luminal side65. Currently, at least four

Ca2+-sensing sites on the RyR2 have been proposed, including two on cytosolic side and

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two on luminal site with different Ca2+ binding affinity at different Ca2+ concentrations66.

(Figure 1.4)

Mg2+ is another important ion that regulates RyR2 activity. Although the cytosolic and SR luminal Mg2+ concentration are not expected to change rapidly during EC coupling66,67, it plays an important regulatory role in EC coupling by regulating RyR2 channel activity in cardiac muscle. Mg2+ could bind to RyR2 and regulate its activity by either binding to less selective inhibitory, low-affinity Ca2+ sites or competing with high- affinity Ca2+ activating sites. Therefore Mg2+ function as a crucial inhibitor of RyR262,68.

ATP is known to regulate the activity of many receptors and channels including

RyR266. Cardiomyocytes usually maintain approximately 3-5 mM ATP67 during the resting condition. Under physiological conditions, ATP predominantly exists as a complex with

Mg2+. Thus, it is proposed that the Mg2+/ATP complex rather than free ATP, regulates

RyR2 in physiological condition.

RyR2 regulatory proteins

Many RyR2 modulators have been identified and extensively studied, including

FKBP12.6, Calmodulin (CaM),39 sorcin,40 protein kinase A (PKA),41 protein phosphatase

1 (PP1), protein phosphatase 2A (PP2A),42 and Ca2+/calmodulin-dependent protein kinase type II (CaMKII).43

FKBPs (FK506-Binding Proteins) are immunophilins that can bind to the FK506 and rapamycin. Immunophilin is peptidylprolyl-cis-trans-isomerase that important in protein folding. Immunophilins are endogenous cytosolic peptidyl-prolyl isomerases that

9

help the correct protein folding. However, FKBPs’ ability to modulate RyR activity is not dependent on the enzymatic activity, instead, they directly interact with RyR to regulate the activity of RyR.

FKBP12.6 is one of the major interaction partners of the channel which binds each

RyR2 subunit69. It has been proposed that FKBP12.6 interacts with an N-terminal site of

RyR2 and inhibits the activity of RyR2 by stabilizing the closed state of RyR2 evidenced by RyR2 at single channel level displayed pronounced subconductance states and enhanced activity70-72. Moreover, FKBP12.6 knockout(KO) mice were more likely to develop heart failure and sudden cardiac death73. Thus FKBP12.6 can prevent Ca2+ leak from the SR and maintain the overall Ca2+ signaling. However, inconsistent results were published from other studies23,74-76 questioning the previous hypothesis. The role of FKBP12.6 in RyR modulation was further challenged by direct measurement of channel activity including single channel recordings, ryanodine binding and Ca2+ imaging77. The results showed that the absence of FKBP12.6 does not affect RyR2 channel function. They also claimed no enhanced susceptibility to stress-induced arrhythmia in FKBP12.6 KO mice which further questioned the hypothesis of FKBP-mediated RyR2 dysfunction. Recent studies have suggested that dual regulation of both FKBP12 and FKBP12.6 may be key for RyR2 channel regulation18 and an alteration in the ratio of FKBP12/FKBP12.6 would contribute to the defective Ca2+ handling seen in heart failure78. Nevertheless, further experiments are needed to determine the underline mechanism for RyR2 regulation by FKBPs.

Calmodulin (CaM) is a 17kDa protein that interacts with RyR2. CaM bind to RyR2 with a high affinity and inhibits the activity of RyR2 channel79. It has been shown that

10

mutations in CaM lead to different cardiac pathological condition such as ventricular tachycardia and sudden cardiac death80-82.

Sorcin is a 21.6 kDa protein that belongs to the penta-EF hand protein family40.

Sorcin is thought to play a role in stabilizing RyR2 both in resting conditions (preventing

Ca2+ leak), but also when RyR2 is open (decreasing amplitude of Ca2+ transients). Since sorcin also binds other proteins involved in Ca2+ signaling, thus further investigation is needed to understand the exact mechanism83.

PKA and CaMKII phosphorylate RyR2 on serine 2808 (Ser2808) and serine 2814

(Ser2814), respectively,41, 43 while PP1 and PP2A dephosphorylate these sites.42 We will later discuss how phosphorylation and dephosphorylation of Ser2808 and Ser2814 affect

RyR2 and SR Ca2+ release. Other proteins interacting with the RyR2 macromolecular complex are junctin and triadin, which anchor RyR2 to the SR membrane,45 and junctophilin-2, which ensures the connection to L-type Ca2+ channels in the plasma membrane.46

RyR2 activity is mainly regulated by phosphorylation of serine and threonine residues47 by PKA41 and CaMKII43, but also by protein kinase C.48 PKA phosphorylates

RyR2 at Ser2808 in small rodents and humans, and Ser2809 in rabbits.49 There has been much focus on the hypothesis stating that hyper-phosphorylation of Ser2808 results in increased RyR2-dependent SR Ca2+ leak in HF.41 Some studies indicate that PKA- dependent phosphorylation increases the open probability of RyR by increasing the sensitivity of Ca2+ dependent activation.38, 41 It has also been suggested that PKA phosphorylation of Ser2808 leads to dissociation of FKBP12.6, destabilizing the RyR2

11

channel and increasing diastolic SR Ca2+ leak.41 However, these effects of PKA-dependent phosphorylation have been chanllenged in later studies.50, 51 Genetic ablation of Ser2808 was not protective against HF,52 and Ser2808 seems to be less involved in the pathogenesis and remodeling process in heart failure than Ser2814.53 In non-failing hypertrophy, however, PKA-dependent phosphorylation of RyR2 has a functional effect on the channel gating.53 This was found in a study where cardiomyocytes from mice with non-failing cardiac hypertrophy showed less Ca2+ sparks when adding a PKA inhibitor.53 The same study showed no changes in Ca2+ spark frequency in cardiomyocytes from mice with HF using the same PKA inhibitor.

RyR2 can be phosphorylated by CaMKII at Ser2814, or serine 2815 in some species.55 In normal physiological situations, without beta adrenoceptor stimulation or pacing, 16 % of Ser2814 residues and 54 % of Ser2808 residues are phosphorylated.56,55

The CaMKII pathway, unlike the PKA pathway, is thought to play an important role in pathological SR Ca2+ leak.57 Knock-in mice with an inactivated Ser2814 CaMKII phosphorylation site in RyR were more resistant than WT mice to developing cardiac fibrosis, pulmonary congestion and cardiac dysfunction following chronic beta adrenoceptor stimulation.58 CaMKII activation and phosphorylation of Ser2814 is also involved in the development of atrial fibrillation in angiotensin-exposed mice,59 while genetic ablation of Ser2814 protected against pacing-induced arrhythmias.60 Interestingly, mice with a constitutive activation of Ser2814 did not develop arrhythmias in basal conditions, without beta adrenoceptor stimulation, but developed VT and SCD in response to beta adrenoceptor stimulation.60 Thus, it seems that CaMKII dependent phosphorylation

12

of RyR2 can play an important role in the development of arrhythmias. CaMKII can be activated by Ca2+, reactive oxygen species (ROS),61 glycosylation,62 and nitrosylation.63

Furthermore, and important for Article 3 in this thesis, high HR also increases CaMKII activity, possibly due to increased intracellular Ca2+ cycling.64

Another mechanism for modulation of RyR2 activity is by oxidation. Production of

ROS is increased by stimulation of beta adrenoceptors and angiotensin II receptors.65,66

Oxidizing conditions increase the open probability of RyR2,67 but increased oxidative stress can also cause irreversible loss of channel activity.68 The FKBP12.6 binding domain on RyR2, which is responsible for keeping the channel closed, can become unstable by increased ROS levels, leading to SR Ca2+ leak.69 Increased oxidation of RyR2 also reduces the binding affinity between CaM and RyR2.69 Interpretation of ROS effects on RyR2 in cellular data is complicated since oxidation also affects the activity of RyR2 modulators and other Ca2+ handling proteins: Oxidation of CaM reduces its binding to RyR2, and oxidation of RyR2 also reduces the binding affinity between RyR2 and CaM.69 Important for Article 2 in this thesis, CaMKII can also be oxidized and activated by ROS, even

70 without increased cytosolic Ca2+ .

SR proteins: calsequestrin, triadin and junctin

The luminal accessory protein calsequestrin (CSQ) acts as the principal Ca2+ buffer in the SR84. CSQ can also directly regulate RyR2 activity by directly interacting with RyR2 with the help of anchoring proteins, triadin and junctin29,65,85. Further, it has been proposed that CSQ/junctin/triadin complex acts as luminal Ca2+ sensor crucial for termination of Ca2+

13

release86. The linkage between mutations in the CSQ gene and ventricular tachycardia further demonstrated the importance of the regulatory role of this complex in pathological conditions87,88.

The need for countercurrents

While many studies have focused on understanding the mechanisms that contribute to control of CICR, the specific details of ionic-flux through the SR/ER during Ca2+ release are still unknown. As discussed previously, during EC-coupling, opening of RyR allows massive amount of Ca2+ moving from the SR to cytosol follow the force of ionic gradient.

However, since Ca2+ is charged cation, this acute efflux of Ca2+ would result in the charge asymmetry across the SR in the form of the accumulation of a negative potential inside the lumen of SR/ER, leading to the inhibition of subsequent Ca2+ release. Likewise, active uptake of Ca2+ into the ER/SR would lead to accumulation of a positive potential in the lumen side of the SR/ER. Thus, without an additional flow of ions to counter the balance of charges, Ca2+ release or uptake during EC coupling would be compromised. Thus, robust counter-ion flux across the SR/ER is crucial to compensate potential change, thus promotes the efficient Ca2+ release and uptake during E-C coupling89-95. While channels selective for monovalent cations have been reported in SR membrane90,94,96-98, searching for the molecular identity of these channels and other accessory proteins that modulate the operation of CICR has emerged as an important area of cardiovascular research.

14

Trimeric Intracellular Cation Channels (TRIC)

Identification of TRIC

Takeshima and colleagues developed an immuno-proteomic method to screen novel proteins that involved in Ca2+ signaling, myogenesis, muscular development and maintenance of membrane integrity in striated muscle99. Multiple approaches including monoclonal antibody-immunohistochemistry, cDNA library screening and gene knockout techniques were utilized systemically. A group of novel proteins were identified and termed mitsugumins (MG). Many studies have been carried out to characterize these proteins since then. Many of these mitsugumins proteins play important roles in muscular and cardiac physiology and diseases. For example, junctophilin (JP), one protein isolated from this immuno-proteomic library, physically links the transverse-tubule to the SR membrane100-104, allowing the formation of triad junctions providing the structural framework for E-C coupling. Recent studies have also linked JP dysfunction and JP polymorphisms to the development of various cardiovascular diseases105-109. MG29 is another protein isolated using the same approach. It is a synaptophysin-related membrane protein that is essential for the maturation and development of the triad structure in skeletal muscle110-112. Interestingly, MG29 may act as a molecular marker of aging that can shield skeletal muscle against aging-related decreases in Ca2+ homeostatic capacity113-116. More recently, another MG protein, MG53, has been shown to be a muscle-specific member of the TRIM family of proteins that plays an important role in the repair of injuries to the plasma membrane of muscle cells117-122. Defects in MG53 function are associated with 15

muscular dystrophy and cardiac dysfunction119,123,124. Encouragingly, recombinant MG53 protein can be used to modulate membrane repair, which would have important implications for the treatment of muscular dystrophy and other membrane-associated human diseases117,125.

In 2007, we discovered TRIC channels located at the SR/ER of multiple cell types126. (Figure 1.5) In the human and mouse genomes, two isoforms of TRIC were identified: TRIC-A is one subtype that is predominantly expressed in the SR of muscle cells while TRIC-B is another subtype ubiquitously expressed in the ER of many different tissues. To study the physiological function of TRIC-A and TRIC-B, we generated Tric-a or Tric-b knockout mice. While Tric-a-/- mice survive normally throughout their lifespan, knocking out TRIC-B in the mice is lethal at the neonatal stage. Moreover, aggravated embryonic lethality is observed with the Tric-a-/-Tric-b-/- mice, demonstrating the essential role of TRIC in development126. Our collaborative studies established that both TRIC-A and TRIC-B are cation channels that are selectively permeable to K+. They also showed a distinct conductance and regulatory properties126,127. We found that genetic ablation of

TRIC-A or TRIC-B lead to compromised gating capacity for K+. Moreover, Ca2+ release across the SR/ER membranewas also affected by absence of TRIC channels, suggesting the hypothesis that the flow of K+ ions into the SR through TRIC during the SR Ca2+ release provided counter-currents that facilitate efficient Ca2+ release126,128,129

Studies from Fill and colleagues showed that the RyR channel could provide certain amount of counter ion movement associated with Ca2+ release from the SR membrane, due to the high permeability of the RyR channel to monovalent cations130-132. Clearly, further

16

studies are required to define the contribution of the intrinsic K+-permeability of the RyR channel and its relationship to the TRIC channel and the overall Ca2+ signaling across the

SR/ER membrane. In the introductory chapter, we summarize key properties of TRIC-A and TRIC-B in controlling intracellular Ca2+ homeostasis and signaling, and provide some recent evidence supporting the role for TRIC in modulating the RyR Ca2+ release channel and operation of store-overload induced Ca2+ release from the SR membrane. These findings highlight the important role of TRIC in cardiac physiology and disease.

Biochemical and biophysical properties of TRIC channels

TRIC-A and TRIC-B contain ~300 amino acids and they share patches of sequence identities and similar hydropathy profiles that suggest the existence of multiple transmembrane segments126 (Figure 1.6). Protein-structural analysis by computer algorithms, originally proposed three or four ER/SR transmembrane segments. The amino and carboxyl-terminal regions are proposed to locate on the SR/ER luminal and cytoplasmic sides respectively based on proteolysis assay of muscle SR vesicles since the amino-terminal region of TRIC-A was resistant to protease digestion, while the carboxyl- terminal tail was sensitive to digestion. The proposed transmembrane topology of TRIC subtypes thus bears an overall resemblance to that of glutamate receptor channels133,134.

Biochemically cross-linking of SR vesicles containing endogenous TRIC-A and purified recombinant TRIC-A protein produced dimeric and trimeric products. Negatively- stained EM imaging of purified TRIC-A shows a homo-trimeric structure126.

17

To characterize the ion channel properties of TRIC-A and TRIC-B, recombinant

TRIC proteins purified from yeast were reconstituted into artificial lipid bilayer membranes to test the channel activity. The lipid bilayer assay showed that both TRIC-A and TRIC-B form displayed cation channel activities that permeable to monovalent cations K+ and Na+ but not to anions and divalent cations127,135. Interestingly, both channels are much more active at positive holding potentials (cytosolic side vs SR luminal side). The channel characteristics observed in the purified TRIC preparations bear close resemblance to the

SR K+ channel previously identified by Christopher Miller and colleagues90,97,136.

Biophysics studies on single TRIC channel show that increasing negative charge in the SR lumen relative to the cytoplasm enhances the channel open probability of TRIC127,135.

Although detailed electrophysiological features remain to be defined, based on current data, we hypothesized that TRIC channels would be ideal counter current carriers to compensate for charge accumulation during SR/ER Ca2+ release and/or Ca2+ uptake.

Role of TRIC-A in SR Ca2+ homeostasis in skeletal and smooth muscles

Skeletal muscle predominantly expresses TRIC-A, with TRIC-A expressed at approximately 10-fold greater level than TRIC-B in skeletal muscle137. Using microsomal membrane vesicles derived from rabbit skeletal muscle, we showed that TRIC-A is abundantly expressed in skeletal muscle, which is ~4-fold higher than that of the RyR (e.g.

TRIC-A/RyR=5, in rabbit skeletal muscle)137. The abundant expression of TRIC-A allows it to be an effective modulator of Ca2+ homeostasis, even if TRIC should be physically excluded to the periphery of the RyR cluster. Ultrastruct ural analysis using electron

18

microscopy revealed the development of vacuolated SR elements with the formation of

Ca2+ deposits in Tric-a-/- skeletal muscle which is rarely observed in wild-type muscle. The

RyR activator caffeine could still be able to release Ca2+ from the overloaded SR in Tric- a-/- muscle, whereas elemental Ca2+ release events, for example, osmotic stress-induced

Ca2+ sparks138, were significantly reduced. Moreover, isolated Tric-a-/- muscle often displayed “alternans” behavior reflected by the transient and alternate increases of contractile force during fatigue stimulations137. Thus, TRIC-A deficiency impairs RyR- mediated Ca2+ release, resulting in SR Ca2+ overload. TRIC-A channels thus probably function as counter-ion channels to support physiological Ca2+ release across the SR in skeletal muscle.

Alternans also occurs in cardiac muscle, but the exact mechanism underlying cardiac alternans is unknown. It has been proposed that altered coupling between RyR- mediated intracellular Ca2+ release may contribute to the alternan phenotype in cardiac muscle21,25,139,140. Furthermore, SR Ca2+ content is thought to be an important determinant of Ca2+ alternans22. Thus, we hypothesize that the instability of the SR Ca2+ release machinery due to overload of the SR Ca2+ store and reduced membrane permeability for

K+ ions lead to the mechanical alternans observed in the Tric-a-/- skeletal muscle.

While the pathological changes that took place in the Tric-a−/− muscle are consistent with a role for TRIC channels in providing counter-ion movements associated with Ca2+ release, it is also possible that TRIC may participate in limiting the electronegative influence of Ca2+ release from the SR, especially under conditions of repetitive stimulations where a succession of fast Ca2+ release events would lead to

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accumulation of negative potential inside the SR lumen. Without the TRIC-mediated

K+ movement, the SR will be more electronegative than normal, which would electrically favor Ca2+ uptake. At the same time, the reduced electrochemical driving force for Ca2+ would work to reduce Ca2+ leak that would promote Ca2+ overload inside the SR. Smaller driving force means smaller Ca2+ currents through the RyR channel, which would make

CICR less likely130-132,141

Tric-a-/- mice develop hypertension even at young-adult stages due to elevated resting tonus of vascular smooth muscle cells (VSMCs)142. There are two Ca2+ release mechanisms known to regulate VSMCs activities: one is local Ca2+ sparks generated by spontaneous RyR opening that activate cell-surface Ca2+ -dependent K+ channels and lead to hyperpolarization; and the other is the agonist-induced IP3 receptor (IP3R) activation that evokes global Ca2+ transients, which could induce contraction of the cells. In Tric-a-/-

VSMCs, RyR-mediated Ca2+ sparks are significantly reduced, which leads to compromised hyperpolarization signaling and thus elevated resting membrane potential. The depolarization promoted the activation of voltage-dependent L-type Ca2+ channels leading to elevated cytosolic Ca2+. As a result, resting myogenic tone of Tric-a-/- VSMCs was enhanced 142.

-/- 2+ In Tric-a condition, RyR mediated Ca release is compromised while IP3Rs function normally, suggesting a differential regulation of RyR and IP3R by TRIC-A channels. Thus, we proposed that TRIC-A preferably supports RyR-mediated Ca2+ -release,

2+ while TRIC-B preferably supports IP3R-mediated Ca release in VSMCs(Figure 1.6). In contrast to the phenotype of Tric-a-/- mice, the transgenic mice overexpressing TRIC-A in

20

smooth muscle develop persistent hypotension which further supports this hypothesis143.

In VSMSs overexpressing TRIC-A, Ca2+ spark is highly elevated and Ca2+-dependent K+ channels are thus activated. Under such hyperpolarizing conditions, the L-type Ca2+ channels are inactivated leading to reduced resting tonus in Tric-a-overexpressing

VSMCs143.

Genetic variations of TRIC and their roles in human diseases

Since TRIC channels are involved in various biological functions as mentioned above, TRIC channels should have played important pathophysiological roles in human diseases. The hypertension phenotype of Tric-a-knockout142 and the hypotension phenotype of Tric-a-transgenic143 suggested that the expression level of TRIC-A channels in VSMCs might determine the resting blood pressure. Several single nucleotide polymorphisms (SNPs) near the TRIC-A gene that increase hypertension risk and diminish the efficacy of antihypertensive drugs have been identified in a Japanese population 142.

We predict that these SNPs might link to a decreased expression level of the TRIC-A in

VSMCs. Therefore, Tric-a SNPs analysis would be a useful biomarker for hypertension.

Contribution of TRIC-A and TRIC-B to Ca2+ signaling in cardiac development and contraction

Through cross-breeding of the Tric-a-/- and Tric-b+/- mice, we found an aggravated embryonic lethality in the TRIC-A and TRIC-B double knockout144 mice, e.g. the DKO embryos die between E9 and E11, perhaps as a result of abnormal heart function. This 21

aggravated lethality suggests that TRIC-A and TRIC-B subtypes share a complementary function in the heart. In the looped cardiac tube from E8.5-9.5 DKO embryos, irregular cytoplasmic vacuoles were formed. EM observations revealed extensively swollen SR/ER structures in DKO myocytes. Such structures were not present in animals that carried a single functional copy of the TRIC-B gene. Fixative solutions contain oxalate, which form electron-dense calcium-oxalate deposits in high Ca2+-containing organelles. Such Ca2+ deposits were frequently detected in the SR/ER in DKO myocytes, but not in Tric-a+/- and

Tric-b+/- DHE (double heterozygotes) myocytes. Fluorometric Ca2+ imaging of cardiac myocytes from the DKO embryos shows that the amplitudes of spontaneous Ca2+ oscillations were depressed at E8.5. However, remarkably larger caffeine-evoked Ca2+ transients were observed in E8.5 DKO myocytes. The elevated caffeine-evoked Ca2+ transients, together with the insoluble deposits in EM observations, indicate severe SR/ER

Ca2+-overloading in DKO myocytes.

In embryonic cardiomyocytes bearing immature intracellular stores, Ca2+ signaling or spontaneous Ca2+ oscillations are predominantly composed of Ca2+ influx, but a significant contribution of RyR2-mediated CICR is also detectable145. RyR2 is the major cardiac Ca2+ release channel that regulates intracellular Ca2+ homeostasis. Lately, results from studies of the inducible, cardiac-specific RyR2 knockdown mice demonstrate that

RyR2 loss-of-function can lead to fatal arrhythmias, which exemplifies the important contribution of RyR2 to cardiac arrhythmia and sudden death in humans16,146. TRIC-DKO and RyR2-knockout mice show cardiac arrest at similar embryonic stages and share similar characteristic phenotypes of swollen and Ca2+-overloaded SR/ER in embryonic cardiac

22

myocytes. These nearly identical phenotypes suggest that RyR2-mediated CICR is diminished in DKO myocytes, despite the fact that the mutant myocytes retained normal expression of major SR Ca2+ store-related proteins. These observations suggest that CICR mediated by RyR2 in Tric-DKO cardiomyocytes is impaired’ and that insufficient RyR2 function leads to SR Ca2+ overload and further disrupts cellular homeostasis in Tric-DKO cardiomyocytes. It would be interesting to know whether this phenotype is linked to altered

Ca2+ signaling in the cardiomyocytes.

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Figure 1.1 EC-coupling

(1) AP travels along sarcolemma. AP activates LTCC on T-tubule leading to Ca2+ influx into the cytoplasm. (3) Influx of Ca2+ activates RyR2 on SR which results in the Ca2+ release from SR. (4) the elevation of cytoplasmic Ca2+ promotes (5) actin-myosin integration and contracting force formation. During diastole, (6) Cytosolic Ca2+ is restored to resting levels by SERCA-mediated pumping of Ca2+ back into SR and Ca2+ extrusion via NCX; (Small panel) the temporal relationship of cardiac AP 147, Ca2+ transient (red) and muscle contraction (blue) during a single ECC event. (Bers 2002).

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Figure 1.2 CICR and SOICR.

The left part of the diagram (in blue) depicts the mechanism of CICR, an action potential activates the voltage-dependent L-type Ca2+ channel, leading to a small Ca2+ influx. This

Ca2+ entry opens the RyR2 channel in the sarcoplasmic reticulum (SR), resulting in SR

Ca2+ release and muscle contraction. The right part of the diagram (in red) denotes the mechanism of SOICR, in which spontaneous SR Ca2+ release or Ca2+ spillover occurs under conditions of SR Ca2+ overload in the stressed condition.

25

Figure 1.3 Structure of RyR2.

A. Top view of closed RyR2. B. Side view of RyR2.

26

Figure 1.4 Ca2+ /Mg2+ regulatory sites on the RyR2.

The hypothetical locations of four divalent cation sites known to regulate the gating activity of RyR2 are shown on a structural. The names given to these sites are indicated on the left and the corresponding Ca2+ affinities of the sites are shown on the right.

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Figure 1.5 Identification of TRIC a, Northern analysis of Tric subtypes b, Western analysis in rabbit skeletal muscle membrane c, Hydropathicity profile of TRIC-A in the Kyte–Doolittle algorithm. M1–M3, putative transmembrane segments. d, Topology model of TRIC-A e, Chemical crosslinking of TRIC-A f, Electron microscopy images of native TRIC-A bound with gold- conjugated Fab fragment (left panel; scale bar, 200 Å). High-magnification images of the immuno-complexes and their contours are also shown (right panels; scale bar, 100 Å). 28

Figure 1.6 Model for TRIC function in Ca2+ signaling

(A) TRIC-A and TRIC-B are two isoforms of the trimeric intracellular cation channels.

Both TRIC-A and TRIC-B channels can conduct monovalent cations to provide the flow of counter currents associated with release of Ca2+ ions from intracellular stores. TRIC-A

2+ modulates RyR-mediated Ca release from the SR, and TRIC-B facilitates IP3R-mediated

Ca2+ release from the ER. Whether or not there is a cross-talk between TRIC-A and TRIC-

B mediated intracellular Ca2+ signaling remains to be explored. Molecular identifies of other channels located on the ER/SR membranes are not known. (B) Topology model of

TRIC channels on the SR/ER membrane. TRIC-A Channel Maintains Store Calcium

Handling by Interacting with Type 2 Ryanodine Receptor in Cardiac Muscle.

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Chapter 2. TRIC-A Channel Maintains Store

Calcium Handling by Interacting with Type 2

Ryanodine Receptor in Cardiac Muscle

Introduction

Ca2+ release from the SR in cardiomyocytes is normally controlled by voltage- dependent Ca2+ influx via the L-type Ca2+ channel through the CICR mechanism14. In addition to CICR, it has long been recognized that SR Ca2+ release can occur spontaneously under conditions of SR Ca2+ overload19-22. Considering its dependence on SR Ca2+ load and independence on membrane depolarization, this spontaneous SR Ca2+ release has been referred to as store-overload-induced Ca2+ release (SOICR)23,24,4.

A number of conditions, including increased extracellular Ca2+ concentrations, high frequency stimulations, excessive beta-adrenergic activation, or digitalis intoxication, can lead to Ca2+ overloading of the SR and subsequently SOICR in cardiac cells20,139. It is also well recognized that SOICR in the form of Ca2+ waves can enhance the activity of the

Na+/Ca2+ exchanger, leading to delayed afterdepolarizations (DADs) and triggered activities3,25,26. These SOICR-evoked DADs and triggered activities are a major cause of ventricular tachyarrhythmias and sudden death in patients with catecholaminergic polymorphic ventricular tachycardia (CPVT) and heart failure3,25. Thus, understanding the

30

molecular basis and regulation of SOICR is critical for the understanding and treatment of

Ca2+-triggered cardiac arrhythmias and other diseases associated with Ca2+ dysregulation.

It is of interest to note that there appears to be no spontaneous Ca2+ waves (SOICR) in cardiomyocytes isolated from the TRIC-A and TRIC-B double knockout mice, despite the heavily overloaded SR. It is unknown why the Ca2+ overloaded SR in DKO cardiomyocytes does not lead to SOICR. It is possible that TRIC is required to control the response of the RyR2 channel to SR luminal Ca2+, and that TRIC-deficiency renders the

RyR2 channel insensitive to luminal Ca2+ activation. Reduced luminal Ca2+ sensitivity of

RyR2 may provide an explanation for the lack of SOICR and the built-up of SR Ca2+ load in the DKO cells. This possible regulation of RyR2 by TRIC in cardiomyocytes has yet to be characterized.

In cardiac muscle, sarcolemma depolarization triggers the release of Ca2+ from the sarcoplasmic reticulum (SR) via Ca2+-induced Ca2+ release (CICR) 14-16. During this process, Ca2+ influx through the voltage-dependent Ca2+ channels on the sarcolemma activates the type 2 ryanodine receptor (RyR2) channel located on the SR. Under certain pathological conditions, spontaneous Ca2+ release from the SR can take place in the absence of membrane excitation due to store overload induced Ca2+ release (SOICR) 4,19-24. SOICR may evoke propagating Ca2+ waves that can further result in delayed after-depolarizations, which may cause arrhythmias in heart failure.

The SR Ca2+ store of cardiomyocytes is maintained by uptake and release processes, both of which are electrogenic events. The release of Ca2+ through the RyR2 channels will lead to the development of a negative potential inside the SR lumen, and this would further

31

limit Ca2+ release from the SR if uncompensated. Likewise, SERCA-mediated uptake of

Ca2+ into the SR would lead to the accumulation of a positive potential within the SR lumen, and that would tend to inhibit Ca2+-pumping function. Thus, coordinated counter- ion movements are required to balance the SR membrane potential in order to maintain efficient Ca2+ release and uptake in cardiomyocytes.

In 2007, Takeshima and colleagues identified trimeric intracellular cation (TRIC) channels located on the SR and endoplasmic reticulum (ER) of multiple cell types1.TRIC-

A and TRIC-B seem to have different functions in Ca2+ signaling in excitable and non- excitable cells99,126,127,129,137,142,143,148-150. The Tric-a-/- mice, in addition to dysfunction of skeletal muscle137, develop hypertension that is linked to defective Ca2+ sparks and spontaneous transient outward currents in arterial smooth muscle142. A common feature with the Tric-a-/- and Tric-b-/- mice is the development of Ca2+ overload inside the SR/ER of multiple tissues126,137,142,151. This Ca2+ overload can impact the function of SOICR, causing instability of Ca+2 movement across the SR membrane in muscle cells, which could further contribute to tissue dysfunctions associated with ablation of the TRIC genes.

While researchers over the past ten years have established that genetic ablations or mutations of TRIC channels are associated with hypertension, muscle dysfunction, respiratory defects, and brittle bone disease2-8, the function of TRIC channels in heart physiology and disease have yet to be established. Recently, the crystal structure of TRIC has been determined, confirming the homo-trimeric structure of a potassium channel152-155.

While the pore architectures of TRIC-A and TRIC-B appear to be conserved, the carboxyl- terminal tail (CTT) domains of TRIC-A and TRIC-B are different from each other. These

32

CTT domains show flexible structure and potentially interfere with crystal formation, thus all available structural determinations of the TRIC channels were obtained without the CTT domain.

In this chapter, we report that cardiomyocytes derived from the Tric-a-/- mice show dysregulated Ca2+ movement across the SR membrane. Our biochemical and immunohistochemical studies show that TRIC-A can physically associate with RyR2 via the CTT domain. Reconstitution studies in lipid bilayer membrane and heterologous cells demonstrate that the functional interaction between TRIC-A and RyR2 can modulate the intracellular Ca2+ homeostasis and consequently the operation of SOICR.

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Methods

Animals

TRIC-A knock out (Tric-a-/-) mice used in this study were reported in our previous publication126. Handling of mice followed IACUC guidelines from the Ohio State

University. Only littermate male mice (10-12 weeks) were included in this study.

Cardiomyocyte isolation and Ca2+ spark measurement

Isolated hearts from adult Tric-a-/- and wild type (WT) littermate mice (10-12 weeks) were perfused with a Langendorff apparatus at 37°C. The enzyme digestion step consisted of perfusing Tyrode's solution containing 1 mg/ml collagenase (Type II, 300

U/mg; Worthington) and 0.1 mg/ml protease (Type XIV) for 6 min. Cardiomyocytes were dissociated from digested ventricles by gentle mechanical dissociation and used immediately after isolation. The Tyrode's solution contained (in mM) 136 NaCl, 5.4 KCl,

0.33 NaH2PO4, 1.0 MgCl2, 10 glucose and 10 HEPES (pH 7.4).

Intracellular Ca2+ spark measurements from isolated mouse cardiomyocytes were conducted using a Zeiss780 confocal microscope with a 40× 1.42 NA oil immersion objective. Cardiomyocytes were loaded with Fluo-4-AM (2 µM) and then stimulated with a field stimulation of 0.5 Hz for 20 seconds in a normal Tyrode's solution containing 1.8 mM Ca2+. Spontaneous Ca2+ spark activities were measured afterward.

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Plasmid construction

The mouse TRIC-A or TRIC-B cDNA was cloned into pCMS-mRFP vector

(Clontech). The pCMS-mRFP plasmid contained a red fluorescent protein (RFP) reporter gene driven under a separate SV40 promoter, allowing for selection of cells transfected with TRIC-A or TRIC-B using red fluorescence. For investigation of the carboxyl-terminal tail (CTT) domain of TRIC on Ca2+ signaling, two chimeras were constructed, TRIC-AB and TRIC-BA. TRIC-AB contains a.a. (amino acid) 1-230 from TRIC-A plus a.a. 229-291 from TRIC-B. TRIC-BA contains a.a. 1-228 from TRIC-B plus a.a. 231-299 from TRIC-

A. The cDNA coding sequences were chemically synthesized (Quintara Biosciences, CA) and cloned into the pCMS-mRFP vector to express the chimeric TRIC proteins in mammalian cells. All plasmid identities were confirmed by gene sequencing.

Single-cell Ca2 + imaging

HEK293 cells with inducible expression of RyR2 were provided by Dr. Wayne

Chen and cultured in DMEM, 5% FBS, 1% penicillin, and streptomycin at 37oC, 5%

CO224. pCMS-mRFP plasmids containing the various TRIC-A and TRIC-B cDNAs were transfected into HEK293 cells using lipofectamine reagent following manufacture’s instruction. 24 hours later, tetracycline (0.1 µg/ml) was added to the culture medium to induce RyR2 expression. Measurements of cytosolic Ca2+ were conducted at 16-18 hours post tetracycline induction. Cells were loaded with Fura-2-AM (5 µM) or Fluo-4 AM (2

µM) in 0 Ca2 + Krebs–Ringer Hepes (KRH) buffer containing (mM): 125 NaCl, 5 KCl,

35

25 HEPES, 6 glucoses, and 1.2 MgCl2, pH 7.4 for 40 min at room temperature. Cells were then continuously perfused with KRH solution containing different concentrations of

CaCl2 (0-2.0 mM) at room temperature. Fura-2 fluorescence was captured by a dual- wavelength excitation spectrofluorometer (Photon Technology International, Monmouth

Junction, NJ) with fast switching of 340 nm and 380nm excitations. Fluorescence of Fluo-

4 was excited at 488 nm and detected by a Zeiss 780 confocal microscope.

ER Ca2+ Measurement

HEK-293 cells with inducible expression of RyR2 were co-transfected with TRIC-

A and D1ER cDNA, or TRIC-B and D1ER to determine the impact of TRIC-A/TRIC-B on the ER Ca2+ stores. Cells were perfused continuously at room temperature with KRH containing 2 mM Ca2+. Tetracaine (2 mM) was used to block RyR2 activity, and caffeine

(10 mM) was used to activate RyR2 to deplete the ER Ca2+ store. Cells were excited at 458 nm, and emissions of yellow fluorescent protein (YFP) and cyan fluorescent protein (CFP) from D1ER were captured every 0.5 s. The amount of fluorescence resonance energy transfer (FRET) was determined by the ratio of the emissions at 535 and 480 nm24,156.

Immunoblotting

HEK293 cells expressing RyR2 and TRIC-A were harvested and lysed in RIPA buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% Triton x-100, 1% sodium deoxycholate, 0.1% SDS, 1 mM EDTA) supplemented with a protease inhibitor cocktail

(PIC, Sigma) for 10 min at 4oC. 10 μg of protein were loaded on 12% or 5% SDS-PAGE 36

and transferred onto a polyvinylidene difluoride membrane for detection of TRIC-A or

RyR2 expressions, respectively. The expression levels of TRIC-A or RyR2 were visualized using specific antibodies anti-RyR2 (34C) (1:3000) (Abcam) and anti-TRIC-A (1:2000)

(Proteintech gr.). Heart lysates derived from wild type mouse were used for positive control of TRIC-A and RyR2.

Immunofluorescence staining

Isolated mouse cardiomyocytes were washed by phosphate buffered saline (PBS) and fixed in 4% phosphate-buffered paraformaldehyde (PFA) for 20 min at room temperature, and then permeabilized for 15 min with 0.2% Triton X-100 in PBS. Cells were blocked by 3% BSA for 40 min and then incubated with TRIC-A and RyR2 antibodies at 4°C overnight. Alexa 647 labeled goat anti-rabbit and Alexa 546 labeled goat anti-mouse secondary antibodies were applied and incubated for 1 h at 37 °C. After a brief wash, DAPI was used to stain the nucleus for 5 min and then washed by PBST 5 times. Images were captured by Zeiss LSM 780 confocal microscope and analyzed by ImageJ.

GST-CTT-A protein expression and beads binding assay

cDNA coding for the CTT domain of TRIC-A (a.a. 231-299) was cloned into the pGEX4T-1 bacteria expression plasmid. The pGEX-CTT-A plasmid was transformed into

E. coli (DH5α), and expression of GST-CTT-A protein was induced with 0.3 mM isopropyl-β-D-thiogalactopyranoside (IPTG, Sigma) for 3.5 h at 30oC. Harvested bacterial pellets were re-suspended in a buffer containing 20 mM Tris-HCl, 200 mM NaCl, and 0.2 37

mM PMSF (pH 7.5) supplemented with 0.5% protease inhibitor cocktails (PIC, Sigma), 1 mg/ml lysozyme, and 1 mg/ml MgCl2, and sonicated (Branson Ultrasonics) for 15 s for 5 cycles at 30% power output. The lysates were centrifuged twice at 15,000 rpm for 30 min at 4oC. The cleared supernatants were diluted 1:4 into the suspension buffer, mixed with

Glutathione Sepharose Beads (Amersham) and incubated at 4oC overnight on an orbital shaker. The mixture was then loaded into a poly-prep chromatography column (Bio-Rad), allowed to precipitate by gravity, and then washed with 50 ml buffer containing 20 mM

Tris-HCl, 200 mM NaCl, 0.2 mM PMSF and PIC, pH 7.5. C1148 cells with stable

157,158 expression of RyR1 were described previously . Microsomal vesicles derived from

C1148 cells or SR vesicles derived from rat heart (kindly provided by Dr. Michael Fill,

Rush University, Chicago) were lysed in RIPA buffer supplemented with PIC and loaded to either GST or GST-CTT-A bound beads and incubated at 4oC on an orbital shaker for 3 hours. The bound proteins were washed 4 times with buffer containing 0.5% NP-40, 20 mM Tris (pH 7.4) supplemented with PIC, eluted with sample loading buffer, and loaded onto SDS-PAGE gel and western blotted with anti-RyR.

Channel activity assay on planar bilayer system

SR vesicles isolated from rat hearts were used for lipid bilayer reconstitution studies of single RyR2 channels159,160. Planar lipid bilayers were formed with a lipid composition of phosphatidylethanolamine and phosphatidylserine (Avanti Polar Lipids, Birmingham,

AL) (1:1 ratio, dissolved in decane, 50 mg/ml) across a 200 μm diameter hole in a polystyrene partition. CTT-A and CTT-B peptides were synthesized by GL Biochem

38

(Shanghai, China) with >95% purity. SR vesicles and CTT-A or CTT-B peptides were added to one chamber using cis design, composed of 250 mM CsCl, 1.1 mM CaCl2, 1 mM

EGTA, 10 mM HEPES, pH 7.4. The other chamber, designated as trans, contained 50 mM

CsCl, 1.1 mM CaCl2, 1 mM EGTA, 10 mM HEPES, pH 7.4. Single channel data acquisition and analysis were conducted with pClamp software.

Microinjection of CTT-A and CTT-B peptide

CTT-A and CTT-B peptides were dissolved in PBS solution at a concentration of

10 µM. Freshly isolated wild type mouse cardiomyocytes were loaded with 5 µM Fluo-4

AM. Microinjection pipettes made of borosilicate glass (Sutter Instrument Co.) were prepared using a micropipette puller (Model P-97, Sutter Instrument Co.). The microfilament inside the pipette ensures that the solution reaches the tip for microinjection.

The tip pore of pipette was adjusted to allow for delivery of 10-20 pico liter of peptide solution in 60 ms using a picospritzer (Parker Instrumentation) coupled to PatchMan micromanipulator (Eppendorf). Changes in intracellular [Ca]i were monitored under a

BioRad confocal microscope.

Statistical analysis

Comparisons between 2 groups were analyzed using Student’s t-test. Comparisons in multiple groups were analyzed with one-way analysis of variance (ANOVA). Statistical significance was denoted by a P value of less than 0.05. All data were presented as mean

± SEM. 39

Results

Abnormal Ca2+ signaling in cardiomyocytes derived from the Tric-a-/- mice

We have generated knockout mice carrying deletion of either TRIC-A or TRIC-B.

While Tric-a-/- mice survived past their adolescent age, homozygous ablation of TRIC-B proved lethal as the Tric-b-/- mice died at the neonatal stage due to respiratory dysfunction151. Moreover, aggravated embryonic fatality was observed with the Tric-a-/-

Tric-b-/- mice, demonstrating the essential role of TRIC in development126,150. Using cardiomyocytes derived from the Tric-a-/- adult mice and littermate wild type (WT) controls (10-12 weeks age), we examined Ca2+ spark signaling properties following electrical pacing. As shown in Figure 2.1A, following a 0.5 Hz electrical field stimulation for 20 s, spontaneous Ca2+ sparks were observed in the isolated cardiomyocytes upon termination of the electric stimulation. Compared with the WT cardiomyocytes, the Tric- a-/- cardiomyocytes showed less frequent Ca2+ sparks (Figure 2.1B, left). Interestingly, the amplitudes of individual Ca2+ sparks appear to be significantly higher in Tric-a-/- cardiomyocytes than those in WT cardiomyocytes (Figure 2.1B, middle).

We also compared caffeine-induced Ca2+ release from the WT and Tric-a-/- cardiomyocytes. As illustrated in (Figure 2.1B, right), the total caffeine-releasable Ca2+ pool from the SR was significantly higher in Tric-a-/- than in WT cardiomyocytes. This

40

observation is consistent with our published data with skeletal muscle8 and epithelial cells7 where the absence of TRIC-A or TRIC-B led to elevated Ca2+ storage inside the SR or ER.

Co-expression of TRIC-A and RyR2 in HEK293 cells modulates SOICR

We used HEK293 cells with tetracycline-inducible expression of RyR2 to investigate the impact of TRIC-A and TRIC-B on RyR2-mediated intracellular Ca2+

23,24 signaling (Figure 2.2A). In this model, elevation of extracellular [Ca]o lead to increased

Ca2+ content inside the ER, which triggered opening of the RyR2 channel via its luminal

Ca2+ sensing mechanism, leading to the appearance of SOICR.

For assaying the effect of TRIC-A in HEK293 cells, we used a dual-reporter plasmid with mRFP-expression cassette driven by a separate promoter. This allows selection of transfected vs non-transfected cells in the same dish (Figure 2.2B). From the line-scan measurements, it is clear that cells co-expressing RyR2 and TRIC-A (red labeled) did not show oscillating patterns of Ca2+ signaling, unlike those cells expressing RyR2 alone, which often display Ca2+ oscillations. A representative trace of Ca2+ measurement in a HEK293 cell with co-expression of TRIC-A and RyR2 (labelled red, Figure 2.2C), illustrates the apparent absence of spontaneous Ca2+ oscillations. After treatment of tetracaine, a RyR2 blocker, SOICR was diminished and cytosolic Ca2+ level was decreased.

At the end of the recording, caffeine was introduced to the dish to test the release of Ca2+ via RyR2. Ca2+ release from TRIC-A transfected cells triggered by caffeine confirmed the apparent suppression of SOICR is not due to the dysfunction of RyR2 (Figure 2.2C, top).

This effect appears to be specific to TRIC-A, as co-expression with TRIC-B did not affect

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RyR2-mediated SOICR in HEK293 cells (Figure 2.2C, bottom). Summary data from multiple experiments is shown in Figure 2.2D, indicating that TRIC-A, not TRIC-B, could modulate SOICR in HEK293 cells.

Studies from Chen and colleagues demonstrated that the frequency of RyR2- mediated SOICR in HEK293 cells was dependent on the concentration of Ca2+ in the extracellular solution24. Here we tested if co-expression of TRIC-A and RyR2 could modulate SOICR in a Ca2+-dependent manner. The data shown in Figure 2.2E clearly suggest that the presence of TRIC-A could alter [Ca]o-dependent activation of SOICR.

Moreover, this effect is TRIC-A specific, since cells with co-expression of TRIC-B and

RyR2 showed a similar response as those transfected with vector alone (as control).

To elucidate the mechanism of TRIC-A mediated modulation of SOICR, we used

D1ER, a Ca2+ biosensor targeted to ER, to directly measure ER luminal Ca2+ level24,156,161.

We followed the approach developed by Bers and colleagues to assay the ER Ca2+ content using tetracaine to inhibit RyR2 activity162,163. This will allow for testing to what extent co- expression of TRIC-A alters the Ca2+ leak rate across the ER. As shown in Figure 2.3A,

HEK293 cells co-expressing RyR2 and TRIC-A displayed reduced ER Ca2+ content

(measured with the D1ER probe), when compared with cells expressing RyR2 alone. While cells expressing RyR2 alone show oscillating patterns of ER Ca2+ content (Figure 2.3A, blue trace) that mirrors the SOICR signal (Figure 2.3B), co-expression with TRIC-A led to apparent suppression of the ER Ca2+ content below the oscillating threshold levels

(Figure 2.3A, red trace). In addition, we observed less Ca2+ release by direct caffeine

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treatment in TRIC-A expressing cells (Figure 2.3B). These data further support that TRIC-

A reduced the ER Ca2+ content in the RyR2 expressing HEK293 cells.

The effect of TRIC-A on ER Ca2+ handling likely requires the participation of the

RyR2 channel, since transfection of TRIC-A in HEK293 cells (in the absence of RyR2)

2+ 2+ did not affect the ER Ca content, as measured by ATP-induced Ca release through IP3R

(Figure 2.3C). Moreover, when blocking ER Ca2+ uptake by cyclopiazonic acid (CPA), the decline of luminal Ca2+ is not affected by TRIC-A. (Figure 2.3D). These results support that altered ER Ca2+ signaling by TRIC-A is RyR2 dependent.

TRIC-A interacts with RyR2 through its carboxyl-terminal tail (CTT) domain

We performed immunohistochemical (IHC) staining of TRIC-A and RyR2 in cardiomyocytes derived from the WT mice. Clear overlap between TRIC-A and RyR2 could be observed (Figure 2.4A). Images were analyzed by Pearson and Manders coefficients and showed significant co-localization (Figure 2.4B).

Topology analysis revealed that both TRIC-A and TRIC-B contained large CTT domains that reside in the cytosol (Figure 2.5A). This CTT domain likely represents a flexible structure that interfered with crystallization and thus was not included in the structural determination of TRIC channels by Liu and colleagues164. The CTTs of TRIC-A and TRIC-B diverge from each other. CTT-A contains a histidine-rich motif and a polylysine domain that are flanked by a hydrophobic domain. This structure is similar to the intracellular loop joining repeats II and III of the L-type Ca2+ channel, which has been

43

shown to be a critical domain that regulates activity of the SR Ca2+ release in muscle cells165-167. Only the polylysine domain appears in the CTT-B domain.

To test if the divergent CTT-A constitutes a site for interaction with the RyR channel, we generated a GST-fusion peptide containing the CTT-A (a.a. 231-299) and

CTT-B (a.a. 229-291). These recombinant peptides were purified from E. coli as GST- fusion proteins. Using a protein pull-down assay, we observed that CTT-A could interact

168,169 with RyR1 stably expressed in CHO cells (Figure 2.5B), and RyR2 from rat heavy

SR vesicles (Figure 2.5C). Using an antibody against RyR2, we could co- immunoprecipitate TRIC-A from HEK293 cells that co-express RyR2 and TRIC-A

(Figure 2.5D). These interactions appear to be specific for CTT-A, as GST peptide alone could not pull down either RyR1 or RyR2. In addition to RyR2, several other candidate proteins were pulled-down by CTT-A as marked by a, b and c in Figure 2.5C. MALDI-

MS identification revealed the a-band contains SERCA, MPP1, and C2CD2; the b-band contains TRIC-A, destrin, Rtn2, and proteolytic fragment derived from RyR2; and the c- band contains mostly a proteolytic fragment for RyR2 and SERCA.

TRIC-A carboxyl-terminal tail peptide increases RyR2 channel activity

Based on these findings, we generated chimeric constructs of TRIC-AB or TRIC-

BA. TRIC-AB contains a.a. 1-230 from TRIC-A plus a.a. 229-291 from TRIC-B. TRIC-

BA contains a.a. 1-228 from TRIC-B plus a.a. 231-299 from TRIC-A. These constructs were co-expressed with RyR2 in HEK293 cells. We found that replacement of the CTT domain in TRIC-A with CTT-B could alleviate the impact on SOICR (Figure 2.6).

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Consistent with the data shown in Figure 2.2, co-expression of TRIC-A with RyR2 could reduce the spontaneous Ca2+ oscillation in HEK293 cells (Figure 2.6A, red color), whereas co-expression of TRIC-B with RyR2 had no effect on SOICR activity (Figure 2.6A, green color). Co-expression of TRIC-AB with RyR2 resulted in elimination of the inhibitory effect of TRIC-A on SOICR (Figure 2.6A, orange color). This finding supports the important function of CTT-A on RyR2-mediated SOICR in HEK293 cells. Interestingly, co-expression of TRIC-BA with RyR2 also had minimal effect on RyR2-mediated SOICR

(Figure 2.6B, blue color), suggesting the possibility that CTT-A alone in the context of

TRIC-B might lose its capability to interact with the RyR2 channel.

We used 10 mM caffeine to induce activation of the RyR2 channel as assessment of the total ER Ca2+ load (Figure 2.6C). Clearly, only TRIC-A had significant impact on the caffeine response, whereas TRIC-B, TRIC-AB and TRIC-BA had negligible effect on

ER Ca2+ content in HEK293 cells expressing RyR2.

For direct evaluation of the CTT-A and CTT-B peptides on RyR2 channel function, we reconstituted the RyR2 channels from rat cardiac muscle into the lipid bilayer membrane. As shown in Figure 2.7A, the addition of CTT-A to the cis-cytoplasmic solution significantly enhanced the RyR2 channel activity (n=5). The effect appears to be specific for CTT-A, as addition of CTT-B peptide did not result in significant changes in

RyR2 channel activity (n=4).

To further evaluate the functional effect of CTT-A and CTT-B peptides on Ca2+ signaling in cardiomyocytes, we performed microinjection of CTT-A or CTT-B peptides into isolated mouse cardiomyocytes (Figure 2.7B). We found that microinjection of CTT-

45

A peptide elicited significantly more intracellular Ca2+ events as compared to those cardiomyocytes injected with CTT-B peptide (Figure 2.7C).

46

Discussion

Overall, our data show that TRIC-A spatially colocalized and physically interact with RyR2 in cardiomyocytes. In addition to the counter-current function to compensate for the accumulation of charges for rapid Ca2+ release of RyR2, TRIC-A also can directly bind to and activate the RyR2 channel activity. In HEK293 with RyR2 expression, introduction of TRIC-A leads to the RyR2 mediated Ca2+ release from ER store in an un- synchronic form. TRIC-A directly modulates RyR2 channel function through its CTT domain.

The present study uncovered a novel function for TRIC-A as an accessory protein that interacts with RyR2 to modulate intracellular Ca2+ signaling. Aside from its recognized role as a counter-ion channel that participates in excitation-contraction coupling of striated muscles, the physiological function of TRIC-A in heart physiology and disease has remained largely unexplored. Even with the recent resolution of the crystal structure of the

TRIC channels, the interacting partners for TRIC have yet to be defined. We showed that

CTT of TRIC-A, an important portion of the channel that was left out of the crystal structure determination, constitutes an active motif that interacts with RyR2 to control

SOICR function. We found that TRIC-B is ineffective in control of RyR2-mediated

SOICR in cultured cells, suggesting the specificity of TRIC-A interaction with the RyR2 channel. Using reconstitution studies, we demonstrated that the synthetic CTT-A peptide could directly enhance the RyR2 channel activity and stimulate intracellular Ca2+ release in isolated cardiomyocytes.

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While many studies have suggested that altered function of SOICR from the SR in cardiomyocytes may contribute to the development of cardiac arrhythmias156,161,170-172,

2+ searching for accessory proteins that modulate RyR2 channel function and SR Ca homeostasis should yield important clues to the function of SOICR in physiological and pathophysiological settings. We know that TRIC-A ablation leads to SR Ca2+ overload, yet the spontaneous Ca2+ sparks are less frequent in cardiomyocytes derived from the Tric-a-/- mice compared with those from the wild type cardiomyocytes. Moreover, such SR Ca2+ overload did not trigger SOICR in the Tric-a-/- cardiomyocytes. Through determination of the ER Ca2+ content, we showed that overexpression of TRIC-A could increase Ca2+ leak across the ER in HEK293 cells. The TRIC-A mediated increase of ER Ca2+ leak reflects direct activation of the RyR2 channel, for expression of TRIC-A alone in the absence of

RyR2 did not affect ER Ca2+ storage or passive Ca2+ leak across the ER. From these studies, we conclude that TRIC-A constitutes a physiological component of the SR Ca2+ release machinery, and TRIC-A deficiency could render RyR2 channels less sensitive to physiological activation of Ca2+ signaling in cardiac muscle.

A recent study from Sitsapesan and colleagues found that in skeletal muscle, the

-/- 2+ RyR1 channels from the Tric-a mice display increased sensitivity to Mg inhibition, defective response to protein kinase A phosphorylation, and physiological activators such

2+ as ATP, and luminal Ca is less effective in activating the individual RyR1 channels reconstituted into the lipid bilayer membrane173. These findings are consistent with a potential role of TRIC-A as an enhancer of RyR channels, such that the absence of TRIC-

A leads to reduced RyR channel function under stress conditions.

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Crystal structures of several TRIC channels from invertebrate and prokaryotic

TRIC channels have been reported152-155. All of these TRIC proteins share very similar folding and trimeric organizations. Interestingly, all of these crystal structures lack the highly flexible CTT domain, which investigators claim may decrease the stability of the crystal152-155. Our findings that TRIC-A directly modulates RyR2-mediated SOICR may have wide implications in cardiovascular research. Potential therapeutic interventions can be used to target the functional interaction between TRIC-A and RyR2 to restore defective

Ca2+ signaling in cardiovascular diseases.

Previous studies from Chen and colleagues showed that reconstitution of RyR2 gain-of-function mutants in HEK293 cells could lead to enhanced SOICR as reflected by increased Ca2+ oscillations and/or reduced threshold of luminal Ca2+ to trigger Ca2+ oscillations23,24,170,174. Interestingly, our study revealed that co-expression of TRIC-A and

RyR2 led to apparent reduction of Ca2+ oscillation in HEK293 cells, which seems unexpected based on the role of TRIC-A as an enhancer of RyR2 function. In fact, when

TRIC-A was introduced into HEK293 cells, increased Ca2+ leak across the ER was observed that may reflect direct activation of the RyR2 channel. It is possible that the

2+ 2+ TRIC-A/RyR2-mediated ER Ca leak may take place in the form of small Ca release events which are un-synchronized and therefore do not appear as Ca2+ oscillations. One would ask why TRIC-A would lead to over activation of RyR2 in HEK293 cells but not in native cardiomyocytes. It is known that coordinated activation and inactivation mechanisms must exist in cardiomyocytes for proper function of Ca2+ signaling in the heart, and many of the stabilizers or inhibitory factors of RyR2-mediaed Ca2+ release are likely

49

absent in HEK293 cells. One avenue of future study may take advantage of HEK293 cells to reconstitute the potential functional interaction of TRIC-A with other regulatory components of the intracellular Ca2+ release machinery.

Our data support the dual function of TRIC-A as a counter-ion channel and an activator of RyR2, both of these functions would enhance RyR2-mediated Ca2+ release in cardiac muscle. A direct interaction between TRIC-A and RyR2 constitutes an important physiologic component of intracellular Ca2+ signaling in the heart. Studies from Fill and colleagues suggested that due to the non-selective nature of the Ca2+ release channels, RyR channels can provide a certain extent of counter currents by themselves130,132,141.

Quantitative simulations suggested that TRIC channels could contribute to the network of

SR membrane potential to support Ca2+ release and reuptake141. Thus, in addition to regulating the acute phase of Ca2+ release from the SR/ER store, TRIC-mediated movement of counter current flow could also play a role in balancing the electronegative influence of

Ca2+ release on other aspects of Ca2+ homeostasis inside the SR/ER.

TRIC-A and TRIC-B have different functions in regulating SR and ER Ca2+ homeostasis in excitable and non-excitable cells, respectively126,137,142,143,149-151. TRIC-B is present in both excitable and non-excitable cells, whereas TRIC-A is predominantly expressed in excitable tissues such as striated muscles and brain tissues. We showed that the divergent CTT domains of TRIC-A and TRIC-B have different functions in modulation of RyR2 channel activity. Previously we have demonstrated that epithelial cells derived from the TRIC-B knockout mice display abnormal function of IP3 receptor (IP3R) mediated

Ca2+ release from the ER store151, and skeletal muscle derived from the TRIC-A knockout

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2+ 137 mice display abnormal function of RyR1-mediated Ca release from the SR store . A

2+ functional crosstalk between IP3R and RyR-mediated Ca signaling has been implicated in muscle and heart cells under physiologic and pathologic conditions175-181. Dissecting the

2+ function of TRIC-A and TRIC-B in modulating RyR/IP3R cross-talk for control of Ca signaling in health and disease will be an important task of future research.

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Figure 2.1 Compromised Ca2+ spark signaling in cardiomyocytes derived from Tric- a-/- mice.

A. Confocal line scan images of Ca2+ sparks in isolated cardiomyocytes from WT and Tric- a-/- mice, after pacing by 0.5 Hz of electrical filed stimulation for 20 s. B. Ca2+ sparks amplitude was increased, and spark frequency was decreased in Tric-a-/- cardiomyocytes, when compared with WT cardiomyocytes. Caffeine-induced Ca2+ release was significantly higher in Tric-a-/- cardiomyocytes (n=6). Data expressed as mean with standard error. *

P<0.05

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Figure 2.2 Effect of TRIC-A and TRIC-B on RyR2-mediated SOICR in HEK293 cells.

A. Western blot of RyR2 and TRIC-A in HEK293 cells with tetracycline-inducible expression of RyR2. Mouse heart lysate was used as control of RyR2 and TRIC-A expression. B. Ca2+ oscillation, representing SOICR, in HEK293 cells expressing RyR2 alone (non-red cell) and lack of Ca2+ oscillation in HEK293 cells co-expressing RyR2 and

TRIC-A (red cell). C. Effect of TRIC-A and TRIC-B on SOICR in cells expressing RyR2 measured with Fura-2. D. Statistical data of percentage of cells with SOICR showed that

SOICR was not affected by TRIC-B, but significantly less with expression of TRIC-A

(***P<0.001) 53

(Continue) Figure 2.2 Effect of TRIC-A and TRIC-B on RyR2-mediated SOICR in

HEK293 cells. E. Co-expression of TRIC-A and RyR2 alters the dependence of SOICR on extracellular Ca2+ concentration. Co-expression of TRIC-B and RyR2 produced similar response of SOICR with changes of [Ca]o, as cells transfected with pCMS-RFP vector control. Data from 4 individual experiments were used for the statistical analyses.

***P<0.001.

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Figure 2.3 Effect of TRIC-A on changes in ER Ca2+ load require interaction with

RyR2.

A. ER luminal Ca2+ measured by D1ER in cells expressing RyR2 (blue), or RyR2 + TRIC-

A (red). 2 mM tetracaine inhibited ER Ca2+ oscillations and increased ER Ca2+ store and

10 mM caffeine caused depletion of ER Ca2+ store. Additional TRIC-A expression reduced

Ca2+ store under 2mM Ca2+ condition compared to RyR2 alone (n=30) B. Cytosolic Ca2+ release by Caffeine indicated reduced Ca2+ store in RyR2 TRIC-A co-expressing cells

(n=27) compared to RyR2 alone (n=110).

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(Continue) Figure 2.3 Effect of TRIC-A on changes in ER Ca2+ load require interaction with RyR2.

C. Cytosolic Ca2+ release by ATP in WT HEK293 (no RyR2) showed same Ca2+ store with or without TRIC-A overexpression. (n=75) D. The D1ER measurement of CPA (SERCA inhibitor) treatment showed that TRIC-A overexpression did not affect the total ER Ca2+ and the ER Ca2+ uptake in WT HEK293 cells. (n=60), *P<0.05 **P<0.01

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Figure 2.4 Co-localization of TRIC-A and RyR2

TRIC-A and RyR2 are highly co-localized together in mouse isolated cardiomyocytes (A).

Images were analyzed by Pearson correlation coefficient and Mander's coefficient (>0.5 indicating significant co-colorization, n=6) (B).

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Figure 2.5 CTT-A interacts with RyR.

A. CTT-A and CTT-B diverge from each other. CTT-A contains a histidine-rich motif

(orange) and a polylysine domain (purple), which are flanked by a hydrophobic domain

(blue). Such structure is similar to the intracellular-loop joining repeats II and III of the L- type Ca2+ channel. Only the polylysine domain is present in CTT-B. B. GST-CTT-A can pull down RyR1 from C1148 cells. C. GST-CTT-A can pull down RyR2 from SR vesicle isolated from rat heart, and three other candidate proteins (labeled as a, b and c.). Western blot confirmed the pull-down of RyR2 from heart SR vesicle. GST alone does not pull down RyR2. D. TRIC-A could be immune-precipitated with RyR2 in HEK293 cells co- expressing RyR2 and TRIC-A.

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Figure 2.6 Chimeras of TRIC-A and TRIC-B show different effects on SOICR in

HEK293 cells.

A. TRIC-AB and TRIC-BA chimeric constructs were used to test the importance of the divergent tail region of TRIC-A. SOICR was measured as previous described. B. SOICR was restored to level of control by TRIC-AB. (n=3) C. ER Ca2+ store also restored by

TRIC-AB. (n=80) ***P<0.001 59

Figure 2.7 CTT-A directly modulate RyR2 channel function.

A. Reconstitution of RyR2 channels in lipid bilayer. Single channel activity was measured with 250 mM Cs (cis)/50 mM Cs (trans) and 1 µM free Ca2+ in the cis solution. Addition of 1 µM CTT-A to cis solution led to significant increase in the bursting pattern of RyR2 channel (n=5), whereas addition of 10 µM TRIC-B-tail did not affect RyR2 activity (n=4).

Statistical analysis indicates CTT-A significantly elevated RyR2 channel open probabilities.

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(Continue) Figure 2.7 CTT-A directly modulate RyR2 channel function.

B. Confocal line scan images of Ca2+ events measured by Fluo-4 after microinjection of

CTT-A or CTT-B into isolated cardiomyocytes. C. CTT-A induced significantly more

Ca2+ release events in cardiomyocytes than CTT-B does (n=11). Data presented as mean with standard error, ***P<0.001

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Chapter 3. TRIC-A Regulates Cross-Talk of Ca2+

Between SR and Mitochondria in Cardiac Muscle

Introduction

Mitochondria play a crucial role for oxidative metabolism that produces 95% energy required for the cell182. Ca2+ is one of the key regulators for mitochondria metabolism. In the matrix, Ca2+ activates Krebs cycle dehydrogenases that regenerate the reduced form of nicotinamide adenine dinucleotide (NADH), donating electrons to the respiratory chain for energy production. Elevated mitochondria Ca2+ would thus accelerate energy production in mitochondria. In the heart, Ca2+ uptake by mitochondria is an important messenger for matching energy demands of the cardiomyocytes through Ca2+- induced activation of Krebs cycle dehydrogenases183,184.

The Krebs cycle is essential not only for ATP production but also to form the H2O2 and ROS (reactive oxygen species) eliminating systems for the mitochondrial matrix and even cytosolic ROS185. Additionally, these mitochondrial antioxidative systems also facilitate the elimination cytosolic ROS186.

An increased metabolic rate would consume more oxygen, resulting in increased respiratory chain electron leakage and ROS production 187. Since Ca2+ primarily promotes

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ATP synthesis by stimulating enzymes of the Krebs cycle and oxidative phosphorylation in the mitochondria, Ca2+ induced elevation of mitochondria metabolic would thus promote

ROS production. Although the role of mitochondrial ROS has not yet been completely understood, it is generally accepted that excessive ROS production by mitochondrial dysfunction may be a prominent feature for aging, cell dysregulation and different diseases188. For a heart, different pathological stressors, such as ischemia/reperfusion and pressure overload, increase the production of mitochondrial ROS which is causally related to progressive heart failure189,190. As underlying mechanisms, ROS activate hypertrophic signaling through oxidation of histone deacetylase 4 and other redox- sensitive pathways191.Furthermore, ROS impair EC coupling by altering the function of

RyR2s, SR Ca2+-ATPase, NCX and other Ca2+ signaling related proteins192-194.

Toxicity of excessive mitochondrial Ca2+ is not only about ROS, but is also involved in promoting apoptosis signaling. Mitochondrial Ca2+ can modulate the mitochondrial permeability transition. Mitochondrial Ca2+ overload induces sustained opening of the mPTP, a pore in the IMM that allows passage of ions and solutes up to 1 kDa. This sudden increase of IMM permeability causes the loss of mitochondria membrane potential and release of proapoptotic factors. The release of proapoptotic factors thus triggers the downstream apoptotic cascade, leading to the apoptotic cell death195.

TRIC-A deficient mice showed altered SR Ca2+ regulation in the heart, for example, the overloading of Ca2+ in SR. However, there are no significant abnormalities in the heart under basal condition in Tric-a-knockout mice. We investigated heart remodeling and SR and mitochondria Ca2+ signaling of cardiomyocytes under the stressed condition such as b-

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adrenergic receptor agonist isoproterenol (Iso) stimulation and the surgical procedure induced stress model: TAC (transverse aortic constriction).

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Methods

Electrocardiography

Powerlab 8/30 ML870 and Animal bioamp ML136 from AD instrument are used to record electrocardiography (ECG) of the experimental animal. 2.5% isoflurane with 1

L/min oxygen was used to induce anesthesia and then adjusted to 1.5% isoflurane during measurement. LabChat 7 was used to analyze the ECG data.

Transverse Aortic Constriction on Mouse

Mice were anesthetized and transferred to a heated platform. Anesthetized mice were intubated, followed by midline cervical incision to expose the aorta. Aortic constriction was achieved by placing a 7.0 nylon suture ligature against a 27-gauge needle on transverse aorta. The needle was removed promptly to create an aortic constriction of

0.4 mm in diameter between the right brachiocephalic and left common carotid arteries. In

Sham operated mice, the procedure was the same other than the constriction of aorta. The mice were maintained on a heating pad for recovery.

Electron microscopy (EM)

Electron microscopy was performed by the Ohio State University Campus

Microscopy and Imaging Facility (CMIF). Heart tissue was sliced in blocks of 1-mm cubes and fixed at room temperature with a fixative solution containing 2.5% glutaraldehyde, 1%

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formaldehyde, and 100 mM sodium phosphate (pH 7.2). These slices were rinsed with 0.1

M Na cacodylate (pH 7.4) and post-fixed for 1 hour in 2% osmium tetroxide. After dehydration, the samples were embedded in epoxy resin, and ultrathin sections (60 nm) were counterstained with uranyl acetate and lead citrate. Transmission electron microscopy images were obtained with a FEI Tecnai G2 Spirit Transmission Electron Microscope (FEI

Hillsboro) equipped with a MacroFire MonoChrome progressive scan charge-coupled device camera (Optronics). Sections of cardiomyocytes and their mitochondria were analyzed.

Cardiomyocyte isolation and Ca2+ spark measurement

Isolated hearts from adult Tric-a-/- and wild type littermate mice (10-12 weeks) were perfused with a Langendorff apparatus at 37°C. The enzyme digestion step consisted of perfusing Tyrode's solution containing 1 mg/ml collagenase (Type II, 300 U/mg;

Worthington) and 0.1 mg/ml protease (Type XIV) for 6 min. Cardiomyocytes were dissociated from digested ventricles by gentle mechanical dissociation and used immediately after isolation. The Tyrode's solution contained (in mM) 136 NaCl, 5.4 KCl,

0.33 NaH2PO4, 1.0 MgCl2, 10 glucose and 10 HEPES (pH 7.4).

Intracellular Ca2+ measurements from isolated mouse cardiomyocytes were conducted using a Zeiss780 confocal microscope with a 40× 1.42 NA oil immersion objective. For Ca2+ measurement, cardiomyocytes were loaded with 2 μM Fluo-4 for 40 min at 37ºC. Fluo-4 dye was excited with 488 nm. For mitochondria Ca2+ measurement,

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cardiomyocytes were loaded with 10um x-rhod-1 for 40min at 37. x-rhod-1 was excited at

594 nm, and its fluorescence was collected at 600–680 nm.

Cardiomyocytes were paced with a 0.5 Hz field stimulation by extracellular platinum electrodes for 20 seconds in a normal Tyrode's solution containing 1.8 mM Ca2+.

Spontaneous Ca2+ spark activities were measured afterward. Data were analyzed by imageJ.

Immunoblotting

Tissues were harvested freshly and lysed in ice cold RIPA buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% Triton x-100, 1% sodium deoxycholate, 0.1% SDS, 1 mM

EDTA) supplemented with a protease inhibitor cocktail (PIC, Sigma) for 10 min at 4oC. 10

μg of protein were loaded on 12% or 5% SDS-PAGE and transferred onto a polyvinylidene difluoride membrane for detection of TRIC-A or RyR2 expressions, respectively. The expression levels of proteins were visualized using specific antibodies anti-TRIC-A

(Proteintech gr, 1:2000), anti-DRP1 (Abcam, 1:2000), anti-OPA1 (Abcam, 1:2000).

Histological analysis

Paraffin-embedded tissue sections of 4 µm thickness were used for hematoxylin and eosin (H&E) staining. Tissue samples were fixed in 4% paraformaldehyde (PFA) overnight at 4°C. After fixation, samples were washed three times for 5 min with 70% ethanol. Washed samples were processed and embedded in paraffin. paraffin sections were cut as slides for pathological staining including H&E, Masson trichrome, and 67

immunofluorescent staining. The tissue blocks were sectioned by Leica microtone into 4-

μm-thick sections on glass slides with a Leica Microtome. To remove paraffin, the slides went through xylene and serial graded ethanol (100%, 95%, 70%, and 50%) and rehydrated in H2O for 5 minutes. Sections were stained with H&E or Masson’s trichrome to detect fibrosis. Quantification of fibrosis was performed using ImageJ.

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Results

Tric-a-/- mice showed abnormal cardiac function and developed cardiac fibrosis under stress conditions.

Previous studies have shown the important role of TRIC-A in Ca2+ regulation in both skeletal and smooth muscle. However, whether TRIC-A plays a role in cardiac physiology and pathology has not been studied. To address this question, we first investigated the cardiac function of wild type (WT) and TRIC-A knock-out mice (Tric-a-/-

). Under normal conditions, Tric-a-/- mice do not show obvious alterations of cardiac morphology or function as compared to those of WT mice. However, after challenged with isoproterenol (Iso) treatment (I.V. Injection 60mg/kg), ECG measurement of Tric-a-/- hearts showed significantly abnormal heart beat events as compared to the normal response in WT mice. (Figure 3.1A) The frequent appearance of irregular RR interval, as well as beats skipping, suggested disrupted cardiac E-C coupling in Tric-a-/- mice under acute stress conditions. (Figure 3.1B) In addition to acute stress, we then tested how those mice would respond to chronic stress. Mice were implanted with a subcutaneous osmotic pump with an Iso release rate of 40mg/kg/day for 14 days to induce chronic cardiac stress.

Pathological data from Masson’s trichrome staining showed significantly increased fibrotic area in Tric-a-/- heart compare to WT (Figure 3.1C) Statistical analysis revealed a significant increase of tissue fibrosis in Tric-a-/- mice under chronic iso treatment. (Figure 69

3.1D) Together, these data indicate that ablation of TRIC-A protein leads to susceptibility of mouse heart to both acute and chronic stress.

Additional ablation of TRIC-B in Tric-a-/- mice showed a much severer cardiac phenotype under normal and stressed conditions.

As a homolog of TRIC-A, TRIC-B is known to be important in Ca2+ regulation. We hypothesized that TRIC-B might play a compensatory role in Tric-a-/- mice. To test this hypothesis, we generated Tric-a-/- Tric-b+/- mice and compared the cardiac function of these mice with that of Tric-a-/- mice. Unlike tric-a single knockout mice, telemetry measurement of ECG of Tric-a-/- Tric-b+/- mice showed that they suffered from bradycardia and AV block even in resting condition, indicating impaired cardiac regulation and function.

(Figure 3.2A) when we performed chronic Iso treatment protocol on Tric-a-/- Tric-b+/- mice

(60mg/kg/day for 14 days). All tested Tric-a-/- Tric-b+/- mice showed progressive bradycardia and sudden death, while all WT and Tric-a -/- mice remained alive. (Figure

3.2B) These results suggested that Tric-a-/- Tric-b+/- mice showed a much severer cardiac dysfunction compared to WT mice. Thus, TRIC-B might play a certain compensatory role for TRIC-A in both basal and stress conditions.

SR Ca2+ signaling is altered in Tric-a-/- neonatal cardiomyocytes

Ca2+ signaling is crucial for cardiac physiology and pathology, and our previous study (summarized in Chapter I) has linked TRIC-A to cardiac RyR regulation of Ca2+ 70

homeostasis. We first tested the Ca2+ signaling of isolated neonatal cardiomyocytes derived from WT and Tric-a-/- pups. Caffeine-induced release of Ca2+ was significantly greater in

Tric-a-/- cardiomyocytes compared to WT cardiomyocytes, indicating Ca2+ overload in

Tric-a-/- SR. (Figure 3.3A). As the line-scan of Ca2+ image showed in Figure 3.3B, Ca2+ sparks frequency was significantly reduced in Tric-a-/- cardiomyocytes compared to that in

WT cells, while amplitude of Ca2+ sparks remained the same between the two groups.

These results indicated that Ca2+ homeostasis was altered in Tric-a-/- mice. Reduced Ca2+ sparks frequency and increased caffeine-induced Ca2+ release in Tric-a-/- cardiomyocytes suggested that lack of TRIC-A might impair the physiological function of RyR2 thus lead to the reduced spontaneous Ca2+ event through opening of RyR2 and consequently SR Ca2+ overload in Tric-a-/- cardiomyocytes.

SR Ca2+ signaling is altered in Tric-a-/- adult cardiomyocytes.

In addition to neonatal cardiomyocytes, adult cardiomyocytes were also isolated for Ca2+ signaling measurement. Synchronized global Ca2+ release via RyR2 was triggered by field stimulation at 0.5 HZ for 20s while Ca2+ waves and sparks were measured by intensity of Fluo-4 dye. (Figure 3.4A) Quantification of line scan Ca2+ imaging showed that the amplitude of Ca2+ transients triggered by field stimulation was higher in Tric-a-/- cardiomyocytes compared to that of WT cells, further supporting previous observation of

SR Ca2+ overload in Tric-a-/- cardiomyocytes. Interestingly, the full width half maximal

(FWHM) of the Ca2+ transients showed a trend towards reduction in Tric-a-/- cardiomyocytes, which might suggest enhanced SERCA pump function in Tric-a-/-

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cardiomyocytes. (Figure 3.4B) Moreover, consistent with neonatal cardiomyocytes, the adults Tric-a-/- cardiomyocytes also showed decreased Ca2+ spark frequency and increased

Ca2+ spark amplitude, suggesting decreased RyR2 activities and overload of SR Ca2+. Since the neonatal Tric-a-/- cardiomyocytes did not show a significant difference in spark amplitude like observed in adult cardiomyocytes, we reasoned that TRIC-A might also play different roles on RyR2 regulation during development.

The heart of Tric-a-/- mice is more susceptible to TAC-induced stress.

Transverse aortic constriction (TAC) is a surgical model frequently used in animals to study pressure overload induced hypertrophy and subsequent heart failure. Since the hearts of Tric-a-/- mice are susceptible to iso-induced stress, we aimed to determine how they would respond to TAC surgery. We first tested the expression dynamics of TRIC-A following TAC injury in WT mice. As it is shown in Figure 3.5A, TRIC-A protein expression level increased significantly at both day 1 and day 2 after the TAC, indicating that TRIC-A would acutely respond to the TAC induced heart stress and damage. At day

2 after TAC, cleaved caspase-3 was significantly increased in Tric-a-/- myocardium while remained unchanged in WT myocardium. (Figure 3.5B) This increased level of cleaved caspase-3 indicated upregulation of apoptosis in the Tric-a-/- myocardium following TAC injury as compared to that in WT myocardium. Eight weeks after TAC, mice hearts were collected for histological analysis. Compared to WT hearts, Tric-a-/- hearts showed a significantly higher level of hypertrophy as evidence by enlarged heart size and ventricular wall thickness (Figure 3.5C). Masson’s trichrome staining also revealed a dramatically

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increased fibrosis area in Tric-a-/- hearts. These data suggest that lack of TRIC-A would impair the physiological response to the stress and result in significantly increased apoptosis of cardiomyocytes. The continuous accumulation of such apoptosis would thus lead to the long-term hypertrophy and fibrosis in the heart.

Mitochondria of Tric-a-/- cardiomyocytes showed more damage under the TAC induced stress

Since the apoptosis of Tric-a-/- myocardium was significantly higher than that of

WT myocardium in response to TAC injury, we first examined mitochondria morphology and functions, as mitochondria plays a key role in activating apoptosis. First, we examined the morphology of the mitochondria by electron microscopy (TEM). It was clear that the mitochondria in Tric-a-/- ventricular cardiomyocytes showed weakened cristae membranes, homogenized matrix, as well as enlarged vacuoles after TAC injury, indicating significant severer mitochondria damage than that in WT cardiomyocytes (Figure 3.6A). Moreover, mitochondria are very dynamic subcellular organelles, they are constantly fissing, fusing and trafficking. These dynamic changes are crucial in stress conditions. During stress, the fusion process of mitochondria is protective and beneficial by integrating the contents of slightly damaged mitochondria as a form of complementation. However, extensive stress would cause the fragmentation of mitochondria via fission process and lead to apoptosis.

DRP1 has been shown to involve in regulating mitochondria fission while OPA1 has been demonstrated as a crucial regulator of mitochondrial fusion, cristae integrity, and mtDNA maintenance. Biochemical analysis showed that Tric-a-/- hearts had significantly reduced

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OPA1 protein levelss compared to WT hearts, while DRP1 level remained unchanged.

(Figure 3.6B) The reduction of OPA1 in Tric-a-/- hearts under stress conditions indicates an impaired protective function of mitochondria that would lead to the disruption of cristae structure, enhanced release of cytochrome c, and the activation of the destructive caspase enzymes, therefore activating execution of apoptosis. However, it was a puzzle for us how alteration of a protein in SR can cause dysfunction of mitochondria and activation of mitochondria mediated apoptosis.

Mitochondria Ca2+ signaling is altered in Tric-a-/- cardiomyocytes via cross-talk to altered SR Ca2+ signaling.

In order to clarify the underlying mechanism, we reasoned that Ca2+ signaling might be a key factor for regulation of mitochondrial integrity and function. To test this hypothesis, we first tested the Ca2+ signaling of mitochondria in cardiomyocytes of TAC injured mice. Fluo-4 AM and x-rhod-1 were used to detect Ca2+ level in cytosolic and mitochondria respectively. (Figure 3.7A) Line-scan images showed that as soon as Ca2+ transient was induced by caffeine in cytosol, mitochondria would spontaneously uptake the

Ca2+ as a result of elevated cytosolic Ca2+. This indicates that the SR Ca2+ release is tightly linked to the mitochondria Ca2+ signaling (Figure 3.7B). Caffeine-induced SR Ca2+ release, as well as mitochondria Ca2+ transients were significantly higher in Tric-a-/- cardiomyocytes as compared to those in WT cardiomyocytes. (Figure 3.7CD) Moreover,

Tric-a-/- cardiomyocytes showed a significantly increased incidence of spontaneous Ca2+ transient compared to that in WT cells after TAC injury. (Figure 3.7E) It is also worth to

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note that although the increase phase of Ca2+ transients were similarly fast in cytosol and mitochondria, the decay rate of Ca2+ transient seemed much slower in mitochondria than that in cytosol (Figure 3.7C). Based on these data, we proposed that TRIC-A deficiency would lead to the elevated SR Ca2+ store which would consequently increase the amount of Ca2+ release during each Ca2+ releasing cycle. Since the amount of Ca2+ uptaken by mitochondria is proportional to the Ca2+ level in cytosol, mitochondria of Tric-a-/- would thus uptake significantly more Ca2+ in response to each Ca2+ release event from SR.

Moreover, the slower Ca2+ clearance of mitochondria would further amplify the upstream imbalance of Ca2+ singling that leads to a more elevated Ca2+ level in mitochondria. Finally, in TAC injury model, Tric-a-/- cardiomyocytes would generate more spontaneous Ca2+ waves than that in WT cells, which would further enhance the Ca2+ overload in mitochondria. Taken together, in Tric-a-/- cardiomyocytes, overload of SR Ca2+ would ultimately lead to the overload of mitochondria Ca2+ that cause mitochondria toxicity.

Mitochondria of Tric-a-/- cardiomyocytes are susceptible to acute oxidative stress while treatment of mitochondria Ca2+ uptake inhibitor RU360 would protect against damage.

If our hypothesis above is correct, one would expect that mitochondria from Tric- a-/- hearts will be easier to injure. To test this, cardiomyocytes isolated from Tric-a-/- and

WT mice were loaded with TMRE for measurement of mitochondria potential in response

-/- to H2O2-induced acute oxidative stress. As compared to WT cells, Tric-a cardiomyocytes showed a significantly greater mitochondria damage as indicated by quicker mitochondria

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potential loss. (Figure 3.8A, top and middle) Interestingly, RU360, an inhibitor of mitochondria Ca2+ uptake, showed a significant protective effect in Tric-a-/- cardiomyocytes treated with H2O2. (Figure 3.8A, bottom) Since the mitochondria form a network in the cells, the loss of mitochondria potential as a result of mitochondria death usually starts from one area of the cells and then quickly propagated across whole cell area.

We observed that this process was relatively fast in Tric-a-/- cardiomyocytes which would lose all mitochondria potential within 2 mins, as compared to 5 mins in WT cells. However, when treated with RU360, this process was significantly prolonged, indicating a protective effect. (Figure 3.8B) In summary, our data showed that ablation of TRIC-A leads to a significantly increased mitochondria damage under the acute oxidative stress, and blocking mitochondria Ca2+ uptake by RU360 would protect the oxidative injury. These results indicate that elevated Ca2+ uptake in oxidative stress condition could be a major contributor to the damage of mitochondria. Inhibiting mitochondria Ca2+ uptake would protect mitochondria against oxidative stress induced injury.

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Discussion

Here we showed the Tric-a-/- mice showed abnormal cardiac function and developed cardiac fibrosis under Iso-induced cardiac stressed condition. Additional ablation of TRIC-B in Tric-a-/- mice showed a much severer cardiac phenotype under both normal and stressed conditions suggesting a compensatory effect of TRIC-A and TRIC-B in the heart. In both neonatal and adult cardiomyocytes of Tric-a-/- mice, SR Ca2+ signaling are altered due to compromised RyR2 activity induced overload of SR Ca2+. In the TAC model, we found the hearts of Tric-a-/- mice is more susceptible to stress due to the increased mitochondria damage as evidenced by EM studies. Mitochondria Ca2+ signaling is altered in Tric-a-/- cardiomyocytes via cross-talk to altered SR Ca2+ signaling. Moreover, mitochondria of Tric-a-/- cardiomyocytes are susceptible to acute oxidative stress while treatment of mitochondria Ca2+ uptake inhibitor RU360 would protect against damage.

Together, our data showed that TRIC-A played an important role in maintaining of SR

Ca2+ homeostasis and the cross-talk to mitochondria Ca2+ homeostasis.

Proposed model that TRIC-A regulates SR Ca2+ homeostasis and the Ca2+ signaling cross-talk between SR and mitochondria.

Based on our experimental observation, we have proposed a working model that

TRIC-A mediates cross talk between SR and mitochondria in basal and stress conditions.

Under normal conditions, TRIC-A contributes to normal Ca2+ homeostasis by promoting normal RyR2 activity and maintaining adequate SR Ca2+ level in WT cardiomyocytes.

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When external stress occurs, many signals would be adjusted in response to such stress.

Specifically, SR Ca2+ signaling would be upregulated during the stress, including the increase of RyR2 activity (Ca2+ release), SERCA activity (Ca2+ uptake), as well as the increase of SR Ca2+ content. These together lead to the increase of Ca2+ transient in each

Ca2+ release events. On one hand, the elevated Ca2+ transient would enhance the force of contraction to accommodate the external stress. On the other hand, elevated Ca2+ release would also increase the uptake of Ca2+ by mitochondria, which serve as the regulatory signal for mitochondria energy production to facilitate the change of other activities.

However, elevated energy production would also enhance the production of ROS that leads to the internal oxidative stress. To fight against elevated endogenous stress, other protective process (like SODs) would also be enhanced. All signaling pathways are tightly linked with each other and well-regulated forming a whole homeostasis process. However, when there is no TRIC-A in cardiomyocytes, SR Ca2+ signaling homeostasis would be impaired.

SR Ca2+ store would be elevated, leading to the increase of SR Ca2+ release. Mitochondria would thus uptake more Ca2+ than it should, leading to overactivated energy production and increased ROS production. Under normal resting condition, this alteration might not cause noticeable problems in the heart. However, during the stressed condition, external stress induced further overload of SR Ca2+, lead to the overload of mitochondria Ca2+ that consequently cause the Ca2+ toxicity to the mitochondria and excessed ROS production. In general, lack of TRIC-A reduced the capacity of accommodation against the stress in the heart, leading to the Ca2+ dysregulation, mitochondria damage, ROS production and

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apoptotic signal activation, leading to the death of cardiomyocytes, and ultimately cause the remodeling of heart morphology and compromised cardiac functionality under stress.

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Figure 3.1 Abnormal heart function in Tric-a-/- mice.

A. ECG of Tric-a-/- mice following injection of Iso (60 mg/kg). B. Variation of RR interval in ECG recording of WT mice (left) and Tric-a-/- mice following Iso treatment. C. Mason’s trichrome staining of heart tissues from Tric-a-/- and WT mice following chronic Iso treatment (40 mg/kg/day) for two weeks. D. Increased tissue fibrosis in Tric-a-/- heart following prolonged Iso treatment (n=15 for wt; 16 for Tric-a-/- mice). Data expressed as mean with standard error. **P<0.01

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Figure 3.2 Iso-induced bradycardia and sudden death in Tric-a-/- Tric-b+/- mice.

A. Representative telemetry ECG recording of mice under resting conditions. Arrows indicate bradycardia and AV block. B. Record of mice heart beats after treatment of Iso

(60 mg/kg/day). The trace with WT mice is representative of 8 other mice tested. All

Tric-a-/- Tric-b+/- mice tested showed progressive bradycardia and sudden death.

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Figure 3.3 Altered Ca2+ signaling in isolated Tric-a-/- neonatal cardiomyocytes.

A. Caffeine-releasable SR Ca2+ content after perfusion of Ca2+, Na-free tyrode solution, in neonatal cardiomyocytes derived from WT (white), Tric-a-/- (red) and Tric-b-/- pups

(blue). *P<0.05. B. The frequency of spontaneous Ca2+ sparks in Tric-a-/- neonatal cardiomyocytes is significantly less than that in WT. **P<0.01. The amplitude of Ca2+ sparks did not change in WT vs Tric-a-/- cells.

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Figure 3.4 Altered Ca2+ signaling in isolated Tric-a-/- adult cardiomyocytes

A. Line-scan confocal image of cardiomyocytes under 0.5 Hz field stimulation. B. Tric-a-

/- cardiomyocytes show increased Ca2+ transient and decreased FWHM during electoral filed stimulation. (n=4 for each) C. Ca2+ sparks frequency was decreased while amplitude was increased in Tric-a-/- cardiomyocytes. (n=17 for Tric-a-/- and 24 for WT *P<0.05)

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Figure 3.5 TAC induced hypertrophy and fibrosis in Tric-a-/- heart.

A. TRIC-A expression increased after TAC surgery in WT heart. B. Cleaved caspase-9 expression level after TAC surgery. (n=3 for each) C. Histology of heart after 8 weeks of

TAC induced stress. D. Mason’s trichrome staining showed increased fibrosis after TAC in Tric-a-/- heart. (n=4 for each, male)

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Figure 3.6 Increased mitochondria damage in Tric-a-/- heart after TAC.

A. EM image of mitochondria of WT and Tric-a-/- left ventricle tissue after 2 weeks of

TAC treatment. More mitochondrial injury (red star) and enlarged vacuoles (purple arrow) could be seen in Tric-a-/- ventricle. (n=3 for each, males) B. WB of mitochondria protein showed decreased OPA1 in Tric-a-/- heart after 2 weeks of TAC. (n=4 for each, male, **P< 0.01)

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Figure 3.7 Altered SR and mitochondria Ca2+ signaling in Tric-a-/- cardiomyocytes after TAC.

A. Isolated cardiomyocytes loaded with fluo-4 and x-rhod-1 for cytosolic and mitochondria Ca2+ measurement respectfully. Representative image (B) and trace (C) of caffeine induced Ca2+ signaling in cytosol and mitochondria. D. Increased cytosolic and mitochondria Ca2+ transient induced by caffeine in Tric-a-/- cardiomyocytes. (n=6 for each, *P<0.05)) E. Increased spontaneous Ca2+ waves in Tric-a-/- (n=8 for each, *P<0.05) 86

Figure 3.8 Acute oxidative stress induces more mitochondria damage in Tric-a-/- cardiomyocytes.

A. Confocal images of cardiomyocytes loaded with TMRE following treatment with

-/- H2O2｡Cardiomyocytes from Tric-a is more susceptible to H2O2 induced loss of

mitochondria potential. B. Striking difference in the time-course of H2O2 induced mitochondrial membrane potential transition with (brown) or without (blue) RU360. (n=2 for each)

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Figure 3.9 Model that TRIC-A regulates SR Ca2+ homeostasis and the Ca2+ signaling cross-talk between SR and mitochondria.

TRIC functions as an accessory protein for RyR to control SR Ca2+ release and crosstalk with mitochondria. Absence of TRIC leads to SR Ca2+ overload and stress-induced Ca2+ toxicity to mitochondria. The dysregulated SR-mitochondria crosstalk contributes to arrhythmia and heart failure.

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Chapter 4. Sustained Elevation of MG53 in the

Bloodstream Increases Tissue Regenerative

Capacity without Compromising Metabolic

Function

During my Ph.D. study, I was also involved in the studies of other biological molecules that play important roles in heart physiology and cardio-protection. I have co- authored 6 papers related to MG53, a cell membrane repair protein196-200. The full list of publications can be found on page vii. I am the co-first author on one manuscript entitled

“Sustained Elevation of MG53 in the Bloodstream Increases Tissue Regenerative Capacity without Compromising Metabolic Function”. Here I summarize the main findings of this paper.

Introduction

Skeletal muscle controls body locomotion and requires an active injury-repair mechanism to maintain its integrity, as contraction-relaxation of muscle fibers often causes injury to the sarcolemma. Developing therapeutic approaches to improve sarcolemma integrity and facilitate regeneration of injured muscle fibers remains a major challenge in muscle physiology research201-203. In addition to being a contractile machine, skeletal

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muscle is recognized as an endocrine organ, secreting a myriad of myokines to modulate other tissue functions204-208. Targeting the myokine function of skeletal muscle and its cross-talk with other tissues is an attractive means to boost the body’s regenerative capacity under physiological and pathological conditions.

MG53 is a member of the TRIM-family protein that serves an essential role in repair of injury to the cell membrane119,209,210. Genetic ablation of MG53 causes compromised membrane repair that leads to progressive skeletal myopathy and decreased regenerative capacity of the cardiomyocytes119,123,209. MG53 is also found in blood circulation at levels correlating with muscle contractile and secretory activities117,196,211,212.

The recombinant human MG53 (rhMG53) protein can protect various cell types against membrane disruption when applied to the extracellular environment and ameliorate pathology associated with muscular dystrophy and lung injury117,199. Intravenous administration of rhMG53 is also effective in protecting against myocardial infarction and acute kidney injury in rodent and large animal models of these diseases198,213. These data support the concept of targeting cell membrane repair in regenerative medicine and present

MG53 as a potential biological reagent for restoration of tissue integrity in a broad range of human diseases.

In addition to cell membrane repair, MG53 contains the conserved RING motif with E3-ligase activity that can participate in regulation of metabolic functions. Song et al214 reported that MG53 protein was increased in animal models of diabetes and proposed that MG53 functions as an E3-ligase to downregulate insulin receptor substrate 1 (IRS-1) serving as a causative factor for the development of diabetes. Yi et al.215 and other

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investigators200,212,216 observed normal MG53 expression in murine models of diabetes and human diabetic muscles. Moreover, early studies with knockout of IRS-1 in mice revealed no clear phenotype of type II diabetes217,218, as the existence of other IRS isoforms may compensate for IRS-1 absence219-222. Therefore, the proposed role for MG53 mediated IRS-

1 downregulation in the manifestation of diabetes lacks a biological base. A recent report by Liu et al showed that cardiac-specific overexpression of MG53 induced cardiomyopathy via transcriptional activation of the peroxisome proliferation-activated receptor

(PPARα)223. While these studies raise concerns over the safety of MG53 overexpression on cardiovascular and metabolic function, no studies have been conducted that investigate the physiological impact of elevating MG53 in blood circulation.

We developed a transgenic mouse model with sustained elevation of MG53 in the bloodstream in order to examine the myokine function of MG53 in tissue injury-repair and regeneration. We observed that the tPA-MG53 mice lived a healthy life-span and displayed enhanced tissue regenerative capacity. Physiological and biochemical studies revealed that sustained elevation of MG53 in the bloodstream did not impact the body’s metabolic function of glucose handling and insulin signaling.

Methods

Experimental Animals

Animal handling and surgical procedures were performed according to protocols approved by the Institutional Animal Care and Use Committee (IACUC) of The Ohio State 91

University and were compliant with guidelines of the American Association for the

Accreditation of Laboratory Animal Care. MG53 knockout mice (mg53-/-) and their wild- type (WT) control mice were bred and maintained as previously described. For generation of the tPA-MG53 mice, human MG53 cDNA was conjugated with a secretory signaling peptide (tPA) at the 5’ end. tPA-MG53 expression construct was generated by cloning tPA-

MG53 into an expression vector containing chicken β-actin promoter. Then the plasmid was sent to the Transgenic/Knock-out Mouse (TG/KO) Facility at Cancer Institute of New

Jersey (CINJ) for transgenic mouse generation. This transgenic mouse line was generated in mixed genetic background of 129/Sv and C57BL/6J and was used for the ear punch experiment (Figure 4.3). Because it was generated earlier than the other tPA-MG53 mouse line, this mouse line was also used for the aging experiment.

A separate tPA-MG53 mouse line was generated from C57BL/6J genetic background by Cyagen Biosciences. Santa Clara, CA, USA. This mouse line was used for all experiments except for the ear punch and aging studies. Both tPA-MG53 mouse lines express high levels of circulating MG53 and developed normally.

Serum MG53 quantification

Two different methods were used to quantify serum MG53 levels in mice. For western blot, different amounts of rhMG53 (0.03, 0.05, 0.1, and 0.3 ng) were mixed with

1 μl of serum from mg53-/- mice to serve as a standard. Serum samples from wild type mouse was loaded with volumes of 1 μl and 2 μl and probed with a monoclonal antibody against MG53. The density of the western blot was plotted against the rhMG53 standard

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concentrations, and regression analysis used to calculate the concentration of MG53 in serum derived from the wild type mice. Similarly, to quantify MG53 in tPA-MG53 mice sera, higher amounts of rhMG53 (1, 5, and 10 ng) were mixed with 1 μl of mg53-/- serum to serve as standard. Each serum sample from tPA-MG53 mouse was loaded with volumes of 0.1 μl, 1 μl, and 2 μl. The density of the western blot was plotted against the rhMG53 protein standard concentration and regression analysis used to calculate the concentration of MG53 in tPA-MG53 serum.

MG53 levels in serum was also quantified with a sandwich ELISA assay. Serum samples derived from tPA-MG53 mice were diluted by 50, 200, and 1000-fold in BSA blocking buffer and loaded to an ELISA plate. To generate standard curve, different amounts of rhMG53 protein spiked into the serum derived from mg53-/- mice, and then further diluted with BSA blocking buffer for 50, 200, and 1000-fold and loaded to the same ELISA plate for quantification. The reading of each standard protein was then plotted against its concentration, and regression analysis used to calculate the concentration of MG53 in tPA-

MG53 serum.

Glucose Tolerance Test

Mice were singly housed and fasted for 16 h prior to tests. The tail tip of the mouse was removed by a sterile scalpel blade. Blood from the tail tip was collected and basal blood glucose was measured by an OneTouch Ultra Glucometer (Lifescan, Milpitas, CA).

Following fasting, mice were intraperitoneally injected with a bolus of glucose (1 g/kg,

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10% D-glucose, freshly prepared in sterile 0.9% NaCl solution). Blood glucose from mouse tail vein was assessed at 15, 30, 45, 90, and 120 min after glucose administration.

Insulin Tolerance Test

Mice were singly housed and fasted for 6 h prior to test. The tail tip of the mouse was removed by a sterile scalpel blade. Blood from the tail tip was collected and basal blood glucose was measured by an OneTouch Ultra Glucometer (Lifescan, Milpitas, CA).

Following fasting, mice were intraperitoneally (i. p.) injected with a bolus of insulin (0.75

U/kg) (Sigma, insulin was freshly prepared in sterile 0.9% NaCl solution). Blood glucose was assessed at 10, 25, 45, and 90 min after insulin administration.

Ear punch injury

A 1-mm ear hole was made in the center of the ear of mice using a metal ear puncher

(Fisher Scientific, Cat. No 50822358). The ear holes were photographed and measured at indicated time points using a caliper. At 14 days after ear punch, the mice were sacrificed, and ears were dissected for histochemical analysis. The paraffin embedded ear samples were cut for trichrome staining to quantify fibrosis of regenerated tissues. For acute injury,

2 hours after ear punch, the ear samples were collected for immunohistochemical analysis of MG53 at the leading edge of ear wound.

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Treadmill running experiment

tPA-MG53 and wild type littermates were initially trained (5 m/min running for 5 mins each time, running for 3 times each day for three days) on a small animal treadmill

(Columbus Instruments). Then the mice were subjected to treadmill running at 10 m/min for 6 hours. Twenty hours after the initial exercise training, mice were subjected to running at 6, 8, 10, 12, 14, and 16 m/min each for 3 minutes on the treadmill to test the capacity of recovery from muscle injury. The number of times the mice fail to run forward and touch the bottom of the electric grid of the treadmill and remain there for over 7 second was recorded as drop-out. Drop-outs of each mouse at each different speed were recorded.

Histopathology and Immunofluorescent Staining

All of the histopathological and immunofluorescent staining were examined in a double-blinded manner. Paraffin-embedded tissue sections of 4 µm thickness were used for hematoxylin and eosin (H&E) staining. Tissue samples were fixed in 4% paraformaldehyde (PFA) overnight at 4°C. After fixation, samples were washed three times for 5 min with 70% ethanol. Washed samples were processed and embedded in paraffin. 4-μm-thick paraffin sections were cut as slides for pathological staining like H&E and Masson trichrome and immunofluorescent staining. Immunofluorescent staining was performed as follows: slides were deparaffinized and rehydrated by incubating successively in xylene, 100% ethanol, 95%, 75%, 50% ethanol and PBS. Antigen retrieval was achieved by heating in the pressure cooker with Tris-EDTA buffer for 13mins. Primary

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antibodies were applied and incubated at 4°C overnight. Goat anti-rabbit/mouse secondary antibody Alexa-546/Alexa-647 were applied and incubated at room temperature for 1 h.

The antibodies used for immunofluorescent staining are: anti-MHC I antibody

(Developmental Studies Hybridoma Bank (DSHB), Iowa), anti-MHC IIa antibody

(DSHB), anti-MHC IIb antibody (DSHB), anti-MHC IIx antibody (DSHB), anti-CD11b antibody (Abcam) and anti-MG53 antibody (homemade monoclonal antibody). DAPI was used to stain the nucleus of the tissue. All images were captured by Zeiss LSM 780 confocal microscope and analyzed by ImageJ.

Immunoblotting

Crude extracts from dissected muscle or heart of experimental animals were washed twice with ice-cold PBS and lysed in RIPA buffer (10 mM Tris-HCl, pH 7.2, 150 mM

NaCl, 1% NP-40, 0.5% SDS, and 0.5% deoxycolate), supplemented with a cocktail of protease inhibitors (Sigma) and phosphatase inhibitors (Thermo Scientific). Heart, muscle lysates or serum samples were separated by 10% SDS-PAGE and transferred onto polyvinylidene fluoride membranes (PVDF) (Millipore). The blots were washed with Tris- buffered saline Tween-20 (TBST), blocked with 5% milk in TBST for 1 hour, and incubated with custom-made monoclonal anti-MG53 antibody198,223 or commercial IRS-1 antibody (Invitrogen), anti-PPARα antibody (NovusBio), anti-IR-β antibody (Cell

Signaling), and anti-Glut-4 (Cell Signaling) antibody. Immunoblots were visualized with an ECL plus kit (Pierce).

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Cardiotoxin induced skeletal muscle injury

At 4-6 months of age, mg53-/-, wild type, and tPA-MG53 mice were treated with 50

µl of 10 μM cardiotoxin VII4 (CTX, Sigma-Aldrich) via intramuscular injection. After 7 days, mice were euthanized, and skeletal muscle were collected for histological analyses.

For all histological analyses, the listed tissues were surgically dissected, fixed in formalin

(Electron Microscopy Services) overnight at 4°C followed by histochemical procedure described above. Analyses were performed double-blinded.

High fat diet feeding

tPA-MG53 and wild type littermates were fed a high fat diet (HFD) (D12079B from

Research Diets, Inc.) for 10 weeks (from age of 6 weeks to 16 weeks). The body weight of the mice was measured every two weeks.

Echocardiography

Echocardiography was performed in mice anesthetized with 2% isoflurane, using a

Vevo2100 echocardiographic system with a 30-MHz transducer (Visualsonics, Inc.,

Toronto, Ontario, Canada). The heart was first viewed using the two-dimensional mode in the parasternal long-axis and then short-axis views. The short-axis views were used to position the M-mode cursor perpendicular to the ventricular septum and LV posterior wall.

LV internal end-diastolic diameters (LVEDD) were measured at the apparent maximal LV diastolic dimensions, and LV internal end-systolic diameters (LVESD) were measured at

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the most anterior systolic excursions of the posterior wall. LV volumes and ejection fractions (EF) were calculated by the Teichholz method. For volumes, LVEDV =

7(LVEDD)3/(2.4 + LVEDD), and LVESV = 7(LVESD)3/(2.4 + LVESD), where LVEDV and LVESV are the LV end diastolic and systolic volumes respectively. For EF, EF (%) =

(LVEDV – LVESV)/LVEDV x 100. LV fractional shortening (FS) was calculated by FS

(%) = (LVEDD - LVESD)/LVEDD x 100.

Statistical Analysis

The data are represented as mean ± SD. Comparisons were made by Student’s t- test when comparing two experimental groups and by ANOVA for repeated measures. A value of P<0.05 was considered significant.

Results

Mice with sustained elevation of MG53 in the bloodstream lived a healthy lifespan

In rodents, MG53 is abundantly expressed in skeletal and cardiac muscle tissues.

Low levels of MG53 can be detected in the bloodstream under normal conditions117,211,212.

To test the myokine function of MG53 in tissue repair and regeneration, we generated a transgenic mouse model with sustained elevation of MG53 in blood circulation (tPA-

MG53) (Figure 4.1). To promote the secretion of MG53 into blood circulation, a tissue plasminogen activator (tPA) sequence was added ahead of the human mg53 cDNA. The

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tPA-MG53 sequence was cloned behind the chicken beta-actin promoter to drive the expression of the transgene.

Western blot analysis showed elevated levels of MG53 protein in sera derived from the tPA-MG53 mice compared to wild type littermates (Figure 4.1a). The enhanced MG53 secretion in the bloodstream of the tPA-MG53 mice was maintained at different ages ranging from 2 months (young), 10-12 months (middle), to 22-24 months (old). On average, the serum level of MG53 in the tPA-MG53 mice was ~120-fold higher than that of the wild type littermates (Figure 4.1b). Quantitative measurement showed that the serum level of MG53 in the tPA-MG53 mice was 5997.1±2071.0 ng/ml (n=9), and the serum level in the wild type mice was 50.4±32.8 ng/ml (n=10). ELISA determination confirmed the elevated MG53 levels in sera derived from the tPA-MG53 mice.

In contrast to the marked elevation of MG53 in the bloodstream, skeletal and cardiac muscles derived from the tPA-MG53 mice showed only a marginal increase over the littermate wild type mice (Figure 4.1c). In addition to the native band of a 53 kDa protein, a higher molecular weight protein above the 53 kDa band was observed in western blot with skeletal muscle derived from the tPA-MG53 mice (Figure 4.1c). This likely represents the tPA-MG53 protein without cleavage of the tPA secretory peptide which has not undergone processing for protein secretion. We also conducted western blot of MG53 expression in non-muscle tissues, e.g. brain, liver, kidney and lung, and found measurable accumulation of MG53 in these tissues.

The tPA-MG53 mice lived a healthy life span with no observable pathology in all major vital organs. Three out of five mice survived over 36 months, whereas all wild type

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control mice died before the age of 30 months. For those tPA-MG53 mice that were sacrificed at 32 months, histological analyses did not reveal obvious pathological changes in any vital organs (Figure 4.1d). Thus, sustained elevation of MG53 in the bloodstream did not have detrimental effects on whole body function. In treadmill testing, the tPA-

MG53 at the age of 32 months could run at a performance level comparable to young wild type mice (6 months).

Previously, we demonstrated that intravenous injection of 1 mg/kg rhMG53 could achieve transient elevation of MG53 in the bloodstream to a level of 10-20 µg/ml, which decays with a half-life time of ~1 hour in the bloodstream198. Such systemic administration of rhMG53 could protect against acute injuries to skeletal muscle, heart, kidney, and lung in animal model studies117,198,199,211,213. The serum level of MG53 in the tPA-MG53 mice

(~2-6 µg/ml) is comparable to the level of exogenous rhMG53 achieved via intravenous administration. This suggests that therapeutic approaches with chronic elevation of MG53 in the bloodstream will likely be safe.

tPA-MG53 mice show normal glucose handling and insulin signaling in skeletal muscle

As a member of the TRIM-family protein, MG53 contains E3-ligase activity which may participate in modulation of metabolic function214,215. Song et al214 reported that the MG53 knockout mice displayed a phenotype of lean body mass under normal diet conditions and were resistant to high-fat-diet (HFD) induced body growth with improved glucose handling after HFD treatment. However, this observation has not been

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recapitulated by other investigators. Specifically, Yi et al215 reported similar body-growth with the wild type and mg53-/- mice following HFD treatment.

With the tPA-MG53 mice, we did not observe any significant difference in their growth pattern from the wild type littermates when subjected to HFD treatment (Figure

4.2a). We used glucose-tolerance test (GTT) and insulin-tolerance test (ITT) to evaluate if tPA-MG53 mice exhibit any alterations in glucose handling. When mice were challenged with a bolus intraperitoneal injection of glucose (1 g/kg), similar glucose handling was observed between tPA-MG53 and wild type littermates at 6 weeks and 30 weeks of age

(Figure 4.2b). Moreover, no significant changes in ITT were observed between wild type and tPA-MG53 mice at 8 weeks and 32 weeks of age (Figure 4.2c). This data suggests that sustained elevation of MG53 in the bloodstream did not have a significant impact on glucose handling.

We next performed western blot analyses of skeletal muscle tissues derived from the wild type and tPA-MG53 mice, focusing on key protein components involved in insulin signaling, e.g. IRS-1, insulin receptor β (IR-β), and Glut-4. Glut-4 is the major membrane transporter that facilitates insulin-mediated glucose uptake into skeletal muscle. As shown in Figure 4.2d, comparable levels of IRS-1 and IR-β were detected in skeletal muscle derived from wild type and tPA-MG53 mice. Furthermore, there was no measurable changes in Glut-4 protein expression in skeletal muscle derived from the tPA-MG53 mice.

These measurements were repeated in multiple experiments (n=6-10 mice, 4-16 months of age) (Figure 4.2e) and observed in both slow twitch and fast twitch skeletal muscles.

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We also did not observe any changes in PPARα expression level in skeletal muscle derived from the tPA-MG53 mice when compared with the wild type muscle. Overall, these data indicate that mice with sustained elevation of MG53 in the blood stream did not show signs of metabolic dysfunction.

The tPA-MG53 mice show increased healing capacity following tissue injury

Previously, we showed that although MG53 is absent from keratinocytes and fibroblasts, remarkable defects in skin architecture were observed in mg53-/- mice, and these animals display delayed wound healing and abnormal scarring197. MG53 present in circulation may contribute to the maintenance of skin architecture under physiological conditions. Here we used an ear punch model, which has been widely used for mammalian wound repair and regeneration224, to assay if increased levels of MG53 in the bloodstream can rejuvenate tissue wound healing capacity.

For this study, a separate tPA-MG53 mouse line with mixed genetic background of

129/Sv and C57BL/6J was used. These mice also have elevated circulating MG53 levels

(Figure 4.3a). A 1-mm through-and-through ear hole was made and monitored for 14 days.

The ear-hole closure was photographed on days 0, 7, and 10. As shown in Figure 4.3b, tPA-MG53 mice show significantly enhanced repair capacity after ear-punch injury as compared to their wild type littermates. The wild type mice did not heal over the 10-day observation whereas the tPA-MG53 mice all healed completely prior to day 10 (Figure

4.3b, 4.3e n=10). More interestingly, staining of the ear tissue with antibody against MG53 revealed concentration of MG53 at the healing edge of the ear hole derived from the tPA-

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MG53 mice (Figure 4.3c), indicating that MG53 from circulation was trafficked to the injury site to facilitate wound closure.

Noticeably, at 10 days post injury, all tPA-MG53 mice recovered completely with no observable scarring. These phenomena were reproducibly observed in multiple injuries to the ears of the same tPA-MG53 mice. As shown in Figure 4.3d, trichrome staining showed clear differences in the ear skin architecture between wild type and tPA-MG53 mice, with both wounds created acutely at 2 hours and 14 days following ear punch.

Quantitative analysis demonstrated significant reduction in fibrosis associated with healing of the ear-punch injury in the tPA-MG53 mice when compared with the littermate wild type mice (Figure 4.3f). From these studies, we conclude that increased levels of MG53 in circulation enhances the tissue wound healing capacity.

Increased muscle performance and injury-regeneration with the tPA-MG53 mice

We conducted experiments to characterize the function of skeletal muscle with the tPA-MG53 mice. Gross anatomy of the soleus, extensor digitorum logus (EDL), and gastrocnemius (Gastro) muscles appears to be comparable between tPA-MG53 and wild type littermates (Figure 4.4a). Quantification of the ratio of muscle weight to tibia length showed no muscle atrophy or hypertrophy in the tPA-MG53 mice (Figure 4.4b). H/E staining of muscle cross section did not reveal detectable pathological changes with the tPA-MG53 muscle. Moreover, cross sectional staining of soleus and EDL muscles with antibodies specific for myosin heavy chain type 1 (MHC1, blue, Figure 4.4c), myosin heavy chain type 2a (MHC2a, green, Figure 4.4c), myosin heavy chain type 2b (MHC2b,

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red, Figure 4.4c) and myosin heavy chain type IIx (MHC2x, purple, Figure 4.4c) did not reveal significant muscle-fiber type switch with the tPA-MG53 mice (Figure 4.4d). The amount of MHC2b, which was present in soleus muscle as a minor component in wild type muscle, was undetectable in the tPA-MG53 soleus (Figure 4.4b).

The tPA-MG53 and littermate wild type mice were subjected to treadmill training to test if elevation of circulating MG53 levels could impact the animals’ running capacity.

For this study, mice were first subjected to running at 10 m/min for 6 hours. Twenty hours after the first round of exercise training, mice were again subjected to running at various speeds for 3 minutes on the treadmill to test the capacity of recovery from exercise induced muscle injury. The number of times the mice fail to run forward and touch the bottom of the electric grid of the treadmill (remaining there for > 7 s) was recorded as drop-out. As shown in Figure 4.4e, the wild type mice had significantly more difficulty on the treadmill as they displayed more drop-outs, in particular at higher running speeds, than the tPA-

MG53 mice. This observation indicates that exercise-induced muscle injury may be less in the tPA-MG53 mice, and their muscles may also recover faster in the presence of sustained elevation of MG53 in blood circulation.

We conducted cardiotoxin-induced muscle injury to evaluate the regenerative capacity of the tPA-MG53 muscle117. Cardiotoxin (50 µl of 10 µM stock solution) was injected intramuscularly into the hind limb muscle of the wild type, mg53-/-, and tPA-MG53 mice. Seven days after cardiotoxin injury, H/E staining was used to evaluate muscle pathology. As shown in Figure 4.5a, there were clear differences between the three muscle samples. The tPA-MG53 muscle showed more homogenous fiber sizes with central nuclei

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(right panel, Figure 4.5a), indicating enhanced muscle regeneration, when compared with those from the wild type (middle panel). The mg53-/- muscle fibers (left panel, Figure 4.5a) showed more heterogeneous fiber sizes. In addition, reduced accumulation of immune cells was clearly visible within the tPA-MG53 muscle at 7 days post cardiotoxin injury. We used

IgG labeling to quantify the extent of muscle necrotic death. As shown in Fig 4.5b, more

IgG positive staining was observed in mg53-/- muscle than in the wild type and tPA-MG53 muscle, which is consistent with the reduced membrane repair function of muscle fibers with ablation of MG53.

To quantify the degree of immune cell infiltration of the injured muscle fibers, we did IHC staining of CD11b. As shown in Figure 4.5c, the tPA-MG53 muscle contained significantly less staining of CD11b than the wild type muscle. The mg53-/- muscle fiber contained the most CD11b staining. Such findings suggest the possibility that mg53 circulation may have a modulatory function to control inflammatory response following muscle injury.

Crossing of tPA-MG53 with db/db mice did not alter insulin signaling nor glucose handling

The tPA-MG53 mice were crossed with db/db mice in order to evaluate if sustained elevation of MG53 in circulation impacts metabolic functions. As shown in Figure 4.6a, the db/db-tPA-MG53 mice displayed similar growth pattern as db/db mice under normal diet condition. Moreover, there were no differences in growth patterns between tPA-MG53 and wild type littermates. We conducted GTT and ITT tests with the four groups of mice:

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wild type, tPA-MG53, db/db, and db/db-tPA-MG53, at the age of 18-20 weeks. As shown in Figure 4.6b, the glucose tolerance of db/db and db/db-tPA-MG53 mice were indistinguishable, as were their insulin tolerance. Consistent with the results shown in Figs.

4.2b and 4.2c, we detected no changes in GTT and ITT between wild type and tPA-MG53 at 18-20 weeks of age.

We used echocardiography to determine if sustained elevation of MG53 in the db/db mice could alter their cardiac function. As shown Figure 4.6c, the ejection fraction and fractional shortening of the heart were similar in db/db-tPA-MG53 and db/db mice at

8-10 weeks of age. Thus, increased MG53 level in circulation did not have any deleterious effects on heart function.

These findings are surprising, as they contradict the previous reports by Song et al214 and Liu et al223. If their hypothesis was correct, we would expect to see drastic exaggeration of the metabolic defects in the db/db-tPA-MG53 mice due to the augmented

MG53 action on IRS-1 and PPARα mediated insulin signaling and lipogenic function. We thus conducted western blot analyses with soleus muscle derived from db/db and db/db- tPA-MG53 mice. As shown in Figure 4.6d, the protein levels for IRS-1, PPARα, and IR-

β did not show clear differences between the two groups.

Liu et al223 showed that mice with cardiac specific overexpression of MG53 led to nuclear translocation of MG53, which can activate PPARα transcription by binding to the

PPARα promoter region. Activation of the PPARα led to lipogenesis in cardiomyocytes, resulting in cardiomyopathy. Interestingly, although we did observe nuclear accumulation

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of MG53 in tPA-MG53 muscle, we failed to observe detectable changes in the PPARα in our mouse lines.

Overall, our findings present the first evidence that elevated MG53 levels in circulation is safe to metabolic and cardiac function.

Discussion

In the present study, we used a transgenic mouse line (tPA-MG53) with sustained elevation of MG53 protein in blood circulation to evaluate the physiological function of

MG53 in tissue injury-repair and regeneration. The tPA-MG53 mice maintain high levels of circulating MG53 throughout their lifespan (up to 36 months) with no observable side effects. In addition to the prolonged lifespan, tPA-MG53 mice display a remarkable regenerative capacity in response to injurious assaults to multiple organs. When challenged with treadmill running and cardiotoxin injury, the tPA-MG53 mice displayed improved exercise performance and enhanced healing ability in response to muscle injury. More excitingly, while the skin tissues do not contain endogenous MG53 protein, ear punch injury to the tPA-MG53 mice can heal remarkably faster than wild type mice. Biochemical and imaging analyses reveal that MG53 can travel in the bloodstream to concentrate on the healing edge of the ear wound sites, thus contributing to the improved regenerative capacity following tissue injury. Together, our data supports the safety of elevated levels of MG53 circulating in our bodies under physiological conditions, and the beneficial effects of

MG53 in the bloodstream on tissue injury-repair and regeneration. Thus, a therapeutic

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approach with intravenous administration of rhMG53 may represent a safe and effective means to treat multiple diseases caused by tissue injuries.

Since our identification of MG53 as a key component of cell membrane repair in

2009, many research groups have joined in this important research area. We know that genetic ablation of MG53 leads to defective cell membrane repair, which can cause progressive skeletal myopathy and decreased regenerative capacity of cardiomyocytes. To evaluate the in vivo functions of MG53, multiple strategies have been employed.

Researchers have used virus based delivery systems to overexpress MG53 in both skeletal muscle125 and hearts123, and found that overexpression of MG53 can protect against injuries to muscle and hearts and ameliorate muscular dystrophy and cardiomyopathy in animal models.

In addition to virus mediated gene delivery, a transgenic approach has also been used by multiple groups to test the physiological function of MG53. Prior to our study with the tPA-MG53 mice, three different lines of MG53 transgenic mice were reported in the literature215,223,225. Xiao and colleagues used both a chicken β-actin promotor215 and α- myosin heavy chain promotor223 to drive MG53 expression in skeletal muscle and the heart respectively. They reported that overexpression of MG53 caused metabolic disorders in mice through down regulation of IRS-1, an E3 ubiquitin substrate of MG53215. With the cardiac-specific MG53 transgenic mice, Liu et al showed that, in addition to IRS-1 down- regulation, MG53 can be found in the nucleus of cardiomyocytes to modulate transcriptional activation of PPARα signaling223, and the elevated PPARα activity could contribute to the hypertrophy and cardiomyopathy observed in their mouse cohort.

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Interestingly, separate studies by Ham and Mahoney reported that cardiac-specific MG53 transgenic mice first showed hypotrophy of the heart at a young age, and only with aging did the transgenic animals begin to develop cardiac hypertrophy225. Contrary to the reduced

IRS-1 level in heart tissues derived from the MG53 transgenic mice reported by Liu et al,

Ham and Mahoney showed elevated IRS-1 expression with their transgenic mice at similar ages225.

Our present study took a different approach to assess the function of MG53 by overexpressing secretory MG53 from striated muscles. Our tPA-MG53 mice contain sustained elevations of MG53 in blood circulation at a level that is close to the therapeutic dose of rhMG53 achieved via intravenous infusion in dogs and pigs198,213. We did not observe any changes in IRS-1 and IR-β in muscle tissues derived from tPA-MG53 mice.

We also failed to detect activation of PPAR-α signaling in both skeletal muscle and heart with our tPA-MG53 mice. The difference between our observations and those of Song et al and Liu et al may lie in the distribution of MG53. It is possible that overexpression of

MG53 inside the muscle and heart tissues may have deleterious effects as reported by Song et al and Liu et al. Our finding suggests that increased levels of MG53 in circulation is beneficial to tissue protection, not harmful to their metabolic function.

We present evidence that sustained elevation of MG53 in circulation did not affect insulin signaling and glucose handling in mice. Moreover, the metabolic and cardiac function of db/db mice were not altered when they were crossed with the tPA-MG53 mice.

These results challenge the suggestion that MG53 functions as a double-edged sword in tissue regeneration and metabolic modulation226. Although some potentially detrimental

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effects from MG53 can take place through intracellular actions on IRS-1 and PPARα signaling pathways, administration of rhMG53 protein through the bloodstream can largely minimize this intracellular action. Due to the relatively short half-life of MG53 protein in blood circulation117,198, it is unlikely the administration of MG53 can impact the body’s metabolic function, especially when using the recombinant MG53 protein to treat acute tissue injuries and dermal wounds.

Given the potential role of MG53 as a myokine in tissue protection, understanding the biological mechanism that regulates secretion of MG53 from muscle cells will be an important task for future researches.

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Figure 4.1 Mouse model with sustained elevation of MG53 in the bloodstream.

(a) 1 µl sera derived from 3-month wild type (WT) and tPA-MG53 mice at 2 month

(young), 12 month (middle) and 24 month (aged) were probed with anti-MG53 antibody.

(b) Quantification of serum levels of MG53 in WT and tPA-MG53 mice by western blot.

(c) Skeletal muscle (2 µg per lane) and heart (2 µg per lane) derived from tPA-MG53 and

WT littermate mice were blotted with anti-MG53 antibody. (d) H&E staining of vital organs from 32-month-old tPA-MG53 mice show normal tissue morphology. 111

Figure 4.2 Assessment of insulin signaling and glucose handling in tPA-MG53 and

WT mice.

(a) tPA-MG53 and WT littermate mice at 6 weeks were treated with HFD and the changes in body weight were followed for 10 weeks (n=5 per group). (b) Glucose tolerance tests were conducted with tPA-MG53 and WT littermates at the age of 6 weeks (left panel) and

30 weeks (right panel). (c) Insulin-tolerance tests were conducted with tPA-MG53 and WT littermates at the age of 8 weeks (left panel) and 32 weeks (right panel). n=6 for WT, n=5 for tPA-MG53. 112

(Continue) Figure 4.2 Assessment of insulin signaling and glucose handling in tPA-

MG53 and WT mice.

(d) TA muscle (60 µg total protein per lane) derived tPA-MG53 and WT littermates were probed with antibodies against IRS-1, IR-β, Glut-4. GAPDH serves as loading control. (e)

Quantification of protein expressions based on western blot.

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Figure 4.3 tPA-MG53 mice show increased healing capacity following ear-punch injury.

(a) tPA-MG53 mice expressed elevated level of circulating MG53 (b) Representative pictures of ear punch injury in WT and tPA-MG53 mice. (c) IHC revealed the MG53 at the leading edge of ear-punch at 2 hrs after wounding in the tPA-MG53 mice. (d) Masson's trichrome staining showed remarkable differences in the ear skin architecture between WT and tPA-MG53 mice, wound at 2 hours (red arrow) and 14 days (yellow rectangle) following ear punch. (e). WT mice do not heal over the 10-day observation, the tPA-MG53 mice all healed completely at day 10. n=10 per group. (f) Quantification revealed significant difference between WT and tPA-MG53 mice 114

Figure 4.4 Increased muscle performance of the tPA-MG53 mice under stress conditions.

(a) Gross anatomy of soleus, EDL and gastrocnemius muscles. (b) no significant differences of muscle weight to tibia length between WT and tPA-MG53 mice. (c) Cross section IHC staining of soleus and EDL skeletal muscle from WT and tPA-MG53 mice for muscle typing. (d) Fiber typing staining results were quantified. (e) 20 hours after the first round of exercise training (10 m/min), mice were again subjected to running at 6,8,10,12,14,16 m/min each for 3 minutes on the treadmill to test the capacity of recovery from muscle injury. The number of drop-outs were quantified. *

P<0.05, ** P<0.01.

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Figure 4.5 tPA-MG53 skeletal muscle showed enhanced regeneration capacity after cardiotoxin injury.

(a) H/E staining of gastrocnemius muscle at 7 days post cardiotoxin injury. (b) IHC staining of MG53 (red) and mouse IgG (green). (c) IHC staining with antibody against

CD11b to show the different degrees of the presence of immune cells at the muscle injury sites. (d) Quantification of muscle fiber size (left), IgG staining (right) and percentage of cells positive for CD11b (right). ***: p<0.001, **: p<0.01. 116

Figure 4.6 Cross of tPA-MG53 mice with db/db mice did not alter the diabetic phenotype.

(a) Growth patterns of WT, tPA-MG53, db/db and db/db-tPA-MG53 littermates under normal diet condition. (b) GTT tests were conducted with the 4 groups of mice at 18 weeks of age (left). ITT tests were conducted at 20 weeks of age. (c) Echocardiogram show similar ejection fraction and fractional shortening between db/db-tPA-MG53 and db/db littermates. (d) Western blot of soleus muscle derived from the 4 groups of mice.

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Chapter 5. Conclusion and Perspective

Summary of findings

To our knowledge, there is no published literature that investigated TRIC-A function in the heart. Moreover, no existing study has been done on TRIC-A regulation of

RyR2 channels. Here we found that TRIC-A is a RyR2 accessary protein that binds and regulates the activity of RyR2 channels, thus it could modulate Ca2+ signaling homeostasis in the cardiomyocytes in order to maintain the normal heart function. Evidence from

HEK293 that overexpression TRIC-A and RyR2 showed the direct functional interaction between these two proteins. TRIC-A promotes the RyR2 activity which reduces Ca2+ overload inside of ER, thus preventing Ca2+ overload-induced Ca2+ release in HEK293 cells. The regulation of RyR2 is mediated through the direct interaction of TRIC-A tail region and RyR2. In sum, in addition to the previous understanding that TRIC-A involves in maintaining a normal level of ER Ca2+ content by promoting counter ion movement supporting efficient RyR2 Ca2+ release, now we found that TRIC-A may directly interact with RyR2 and regulate its activity in response to store/luminal Ca2+, thus controlling the occurrence of SOICR.

Many studies have suggested that altered function of SOICR from the SR in cardiomyocytes may contribute to the development of cardiac arrhythmias. For example, overload of Ca2+ in SR leads to store overload induced Ca2+ release and causes the uncontrolled DADs and triggered activities. These SOICR-evoked DADs and triggered

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activities are a major cause of ventricular tachyarrhythmias and sudden death in patients with catecholaminergic polymorphic ventricular tachycardia (CPVT) and heart failure3,25.

Thus, understanding the molecular basis and regulation of SOICR is critical for the understanding and treatment of Ca2+-triggered cardiac arrhythmias and other diseases associated with Ca2+ dysregulation. Thus, searching for accessory proteins that modulate the RyR2 channel function and SR Ca2+ homeostasis should yield important clues to the function of SOICR in physiological and pathophysiological settings. Our finding that

TRIC-A can potentially modulate RyR2-mediated SOICR, could lead to development of potential therapeutic interventions to target the TRIC/RyR interaction for restoring defective Ca2+signaling in cardiovascular and potentially other human diseases.

The discovery of TRIC channels has potential importance for our understanding of

Ca2+ signaling and homeostasis in the heart and other tissues. One question that requires further investigation is the extent to which TRIC contributes to counter ion movement. In addition to TRIC, additional molecules are also likely to be involved in balancing the ion fluxes across the SR/ER membrane. For example, high H+ permeability is detected in the

SR/ER membrane and is, in part, responsible for the counter-transport of H+ and Ca2+ mediated by Ca2+ pumps227. Along with the SR K+ channel, several other K+ and Cl−- selective currents were detected in intracellular organelles, whereas their molecular identities remain to be solved. Recent studies from Fill and colleagues suggested that RyR and IP3R channels can provide certain extent of counter-current movement due to the non- selective nature of the Ca2+ release channels. In their recent publication, Guo et al130 used pharmacological inhibitors of the SR K+ channel and concluded that TRIC-mediated

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counter ion movement does not contribute to the overall SR Ca2+ release property in cardiac muscle. Note that their experiments were conducted using the replacement of K+ ions with

Cs+ ions that resulted in only ~70% inhibition of the cation current through the SR K+ channel, which may not be sufficient to cause detectable impact on the SR Ca2+ release property. Direct evaluation of the role of both TRIC channel subtypes in Ca2+ signaling will require the use of specific and potent pharmacological inhibitors that can produce complete inhibition of the TRIC channels. Alternatively, tissue specific manipulation of

TRIC-A or TRIC-B expression in transgenic mice may provide useful models to examine the physiological function of these channels in cardiovascular physiology or disease. For overcoming the lethality of the germline ablation of TRIC-B and TRIC-A genes, inducible or targeted siRNA silencing of both TRIC-A and TRIC-B may be required in order to define the physiological function of TRIC subtypes in adult muscle and heart cells.

It is clear that TRIC-A and TRIC-B have differential functions in regulating SR and

ER Ca2+ homeostasis. TRIC-B is ubiquitously expressed in all tissues and, considering the lethal phenotype produced by TRIC-B ablation, one can envision that TRIC-B may play an essential role in maintaining normal cellular function in a wide variety of cell types.

TRIC-A expression is predominantly targeted to tissues containing excitable cell types, such as the brain and muscles, suggesting that TRIC-A may function to meet particular kinetic demands of Ca2+ release within excitable cells. A reduction in TRIC protein level would likely lead to instability in the ER/SR Ca2+ release process, which would have wide reaching effects in cellular physiology. It is speculative to hypothesize that TRIC-A may interact with the RyR channel and TRIC-B may associate with the IP3R channel to

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modulate the overall SR and ER Ca2+ homeostasis. While many studies have shown that

2+ cross-talk between IP3R and RyR can modulate Ca signaling in muscle and heart cells in response to physiological and pathological stresses175-181, the potential role of TRIC-A and

TRIC-B in mediating IP3R/RyR cross-talk for regulation of Ca2+ signaling is an exciting area of future research.

Several lines of evidence have linked mutations in TRIC-B to bone and pulmonary diseases and mutations in TRIC-A to hypertension and muscular diseases137,142,143,149,151,228,229. Expansion of these research efforts should provide new insights into the physiological function of TRIC channels in human health and disease. For example, one area of cardiovascular research may focus on establishing the link of genetic mutations in TRIC-A or TRIC-B to the development of arrhythmia and other stress- induced heart diseases, and whether these are correlated with the altered intracellular Ca2+ signaling and homeostasis in the cardiovascular system.

Mitochondria also play an important role in cardiomyocytes function by providing sufficient energy for normal heart function. As mentioned previously, mitochondria Ca2+ is an important regulator of mitochondrial function as well as a potential player in the development of heart failure. Since the SR and mitochondria are in close proximity, it is speculated that SR Ca2+ dysregulation would cause mitochondria disfunction through cross-talk of Ca2+ signal. Our study of Tric-a-/- mice cardiomyocytes provides evidence that compromised SR Ca2+ homeostasis indeed alters mitochondria dysfunction through crosstalk of Ca2+ signaling and consequently leads to cardiac remolding and impaired heart function during stressed condition. Absence of TRIC-A alters the normal SR Ca2+

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homeostasis which cross-talks to and affects mitochondria Ca2+ hemostasis. The Ca2+ dysregulation of mitochondria in the heart reduces the capacity of mitochondria to response to the stress, leading to mitochondria damage, ROS overproduction and apoptotic signal activation under stressed condition. All these factors together cause the death of cardiomyocytes, and ultimately lead to the remodeling of heart such as hypertrophy and heart failure. Therefore, targeting TRIC-A and the cross-talk of SR/ER-mitochondria Ca2+ homeostasis represents a novel therapeutic approach in prevention and treatment of cardiovascular disease. Further studies are needed for development of translational research.

Future work

Although our studies have provided new and crucial understanding of how TRIC-

A modulates RyR2 activity and Ca2+ signaling, there are some limitations in our studies that could be further explored and expanded in the future.

First, although HEK293 cells provided a simple model that is easy to perform and analyze the direct effect of RyR2 mediated SOICR, these cells cannot fully represent the real condition in cardiomyocytes and in the heart since they lack the full sets of RyR2 regulatory machinery, including activation and inhibition regulators. More elegant studies performed under physiological conditions such as cardiomyocytes and intact heart are needed to confirm and explore the detailed mechanism of RyR2 regulation by TRIC-A. To achieve this, we could perform gene rescue or silencing studies to test if acute manipulation

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of TRIC-A would have any effects on Ca2+ signaling in cardiac muscle. We could deliver shRNA probe targeting TRIC-A into neonatal CMs using AAV-mediated gene delivery.

Second, at present, we do not know if TRIC-A interacts with RyR2 more in the closed or open state of the Ca2+ release channel. Using lipid bilayer single channel measurements, we can derive if the TRIC-A-tail acts on the open or closed state of the

RyR2 channel through analysis of the dose-dependent effect of the peptide on the open and closed lifetimes of the Ca2+ release channel. [3H]ryanodine binding assay would also be valuable to determine if TRIC-A-tail peptide alters the Kd or Bmax of ryanodine binding to

RyR2 in heavy SR vesicles derived from the WT mice. Moreover, comparison with

3 -/- [ H]ryanodine binding to Tric-a heavy SR vesicles can provide information on whether

TRIC-A favors open or closed state of RyR2 channel.

Third, although we have had certain understanding that TRIC-A and TRIC-B are both important in Ca2+ signaling regulation, detailed mechanism of how they work together or how they compensate for each other remains largely unknown. Since Tric-b-/- is lethal, our current approach initially just used Tric-a-/-Tric-b+/- mice for the functional study.

However, understanding how they function in the existence and absence with each other acutely would be more useful in understanding their compensatory role in cardiac physiology and pathology. In order to overcome this limitation, acute gene manipulation of TRIC-A and TRIC-B by AAV-mediated gene silencing or gene rescue could be our next approach to elucidate the functional relationship of TRIC-A and TRIC-B. For example, in the TRIC-B-/- neonatal CMs, we could use shRNA to silence the expression of TRIC-A to test if the absence of both TRIC-A and TRIC-B can have additional effects on the Ca2+

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signaling crosstalk from SR to mitochondria. On the other hand, we could test if overexpression of TRIC-A globally in the Tric-b-/- neonatal mice could compensate the absence of TRIC-B and rescue the lethal phenotype of Tric-b-/-.

Transgenic mice with controlled expression of TRIC-A/B would be another very useful in vivo model to study their roles Ca2+ signaling and progression of heart failure.

For establishing transgenic animal models for cardiovascular research, we recently developed a novel transgenic system for inducible and tissue specific regulation of gene expression in mouse models. The system employs a tetracycline-responsive CMV promoter that controls transcription of a cDNA or a small-hairpin (sh) RNA against the target gene that remains non-functional until an interrupting reporter gene cassette is excised by Cre recombinasesue or lineage-specific expression of siRNA (or cDNA) is achieved with inducer mice with tissue-specific expression of Cre230. This transgenic approach could be applied for inducible and reversible control of TRIC genes expression in the heart and other mouse tissues, thus providing a versatile system for elucidating the physiological function for TRIC using viable animal models.

Fourth, although the use of RU360 provides certain clues about the role of Ca2+ entry into mitochondria and its impact on mitochondrial integrity, the underlying molecular insight into the role of mitochondrial Ca2+ uptake in controlling stress response of the CMs is still not very clear. To address this question, we could use CRISPR-gene silencing of

MCU in CMs to examine if knockout of MCU1 can prevent SR-Ca overload induced

2+ -/- mitochondrial Ca toxicity in Tric-a CMs. More studies on how exactly TRIC-A regulates mitochondria Ca2+ are needed.

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Fifth, it is still not very clear how mutations in TRIC-A contribute to human cardiac disease. Interestingly, several human SNPs around TRIC-A have been identified that link to human cardiac function. K107N, is among one of such mutations identified in human patients with stress-induced arrhythmia. According to resolved crystal structure of TRIC channel152, K107 is located in the pore region of the channel. Studies with K107N will be interesting to show that, besides the TRIC-A-tail domain, channel gating domain still represents an important role cardiac physiology and pathology. Thus, it would be important to introduce K107N mutation into mouse models, and test whether this mutation alters K- permeability across the SR or the interaction with RyR2 as a causative factor for progression of heart dysfunction associated with stresses.

As mentioned previously, we have shown evidence of altered Ca2+ cross-talk between SR and mitochondria in Tric-a-/- mice cardiomyocytes which lead to compromised resistance to Ca2+ overload and Ca2+ toxicity in mitochondria under the stressed condition.

The prolonged damage of mitochondria would lead to the apoptosis of the cardiomyocytes and cell death. The death of cardiomyocytes would trigger inflammation and eventually develop fibrosis in the heart tissue and remodeling of the heart such as hypertrophy.

Although this hypothesis seems very plausible based on our results, we did not rule out the possibility of other TRIC-A associated pathways playing a role in the fibrosis and hypertrophy of the heart. One possibility is that TRIC-A might play a role in translational regulation of genes in the nucleus.

Recently, we have observed noticeable expression of TRIC-A on nuclear envelope of cardiomyocytes (Figure 5.1). It has been observed that cardiomyocytes develop

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membrane invagination of nuclear envelope. Researchers speculated that the invagination would play a role in gene regulation231. However detailed information is largely lacking.

TRIC-A located directly on the invaginating membrane structure of nuclear envelope suggesting it might play a role in nuclear Ca2+ regulation and gene translational regulation.

Moreover, in TAC model, we found that the nuclear envelope invagination is significantly impaired in Tric-a-/- cardiomyocytes compared to WT (Figure 5.2) suggesting TRIC-A might also help maintain the functional structure of nuclear envelop invagination. Further studies are required for understanding the exact role that TRIC-A plays in the nuclear envelope of cardiomyocytes.

Another interesting finding comes from the immunofluorescent staining of TRIC-

A in the brain tissue of the mouse. We have observed a very strong expression of TRIC-A in the neuronal cells in the brains (Figure 5.3). Purkinje cells and neurons located on hippocampus are among the cells with highest expression of TRIC-A. Since ER and mitochondria Ca2+ signaling also play crucial roles in neuronal cells, it would not be surprising that TRIC-A would play a role in neuron cell excitation activity and stress response. Further studies are needed to explore the function of TRIC-A in the brain.

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Figure 5.1 Immunofluorescent staining showed TRIC-A is expressed on the nuclear envelope of cardiomyocytes.

Left: Co-staining of TRIC-A (red) and lamin A (green) and Dapi (blue) showed colocalization of TRIC-A on nuclear envelope. Right top: TRIC-A expressed on invagination structure of nuclear envelope in cardiomyocytes. Right bottom: Chromotins are condensed close to TRIC-A enriched membrane invagination.

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Figure 5.2 TAC significantly reduced invagination of nuclear envelope in Tric-a-/- cardiomyocytes.

Top: After TAC surgery, membrane invagination of nuclear envelop in Tric-a-/- cardiomyocytes is significantly less than in WT. Bottom: Statistical analysis revealed a significant reduce of nuclear envelope invagination.

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Figure 5.3 Immunofluorescent staining showed TRIC-A is highly expressed in mouse brain.

TRIC-A is expressed in different excitable neuronal cells such as Purkinje nerve cells, and neurons in hippocampus area.

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