Wayne State University

Wayne State University Dissertations

1-1-2014 Junctional Sarcoplasmic Reticulum Processing And Trafficking In Cardiac Tissue And Primary Cultured Cardiomyocytes Naama Sleiman Wayne State University,

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Recommended Citation Sleiman, Naama, "Junctional Sarcoplasmic Reticulum Protein Processing And Trafficking In Cardiac Tissue And Primary Cultured Cardiomyocytes" (2014). Wayne State University Dissertations. Paper 1056.

This Open Access Dissertation is brought to you for free and open access by DigitalCommons@WayneState. It has been accepted for inclusion in Wayne State University Dissertations by an authorized administrator of DigitalCommons@WayneState. JUNCTIONAL SARCOPLASMIC RETICULUM PROTEIN PROCESSING AND TRAFFICKING IN CARDIAC TISSUE AND PRIMARY CULTURED CARDIOMYOCYTES

by

NA’AMA H. SLEIMAN

DISSERTATION

Submitted to the Graduate School

of Wayne State University,

Detroit, Michigan

in partial fulfillment of the requirements

for the degree of

DOCTOR OF PHILOSOPHY

2014

MAJOR: PHYSIOLOGY

Approved by:

______Advisor Date

______

______

______

______

© COPYRIGHT BY

NA’AMA H. SLEIMAN

2014

All Rights Reserved

DEDICATION

I dedicate this work to my dear Parents Hassan and Sakna, for their endless love and support throughout my life. To my Uncle, for his encouragement through my early college years. To my Daughter, Serena whom was and is my angel, source of happiness and inspiration as I watched her grow and learn during my Doctoral pursuit and will always encourage and wish to see accomplish success in her future endeavors.

To the advancement of science and medicine.

ii

ACKNOWLEDGEMENTS

A deep appreciation to my Dissertation advisor, Steven Cala, Ph.D. whom I admire for his passion to science and his work, for his guidance, expertise, countless hours of support and most of all his patience throughout the process. Without whom this completion would not have been possible. I would like to individually thank the members of my dissertation committee Jian-Ping Jin M.D., Ph.D., Noreen Rossi, M.D.,

Assia Shisheva Ph.D., Todd Leff, Ph.D for their supervision and goodwill. I extend my gratitude to the Department of Physiology Graduate Officer Douglas Yingst, Ph.D. for his support. Ms. Christine Cupps for her remarkable assistance and continuous encouragement. I am very grateful to the entire Faculty and staff in the Physiology

Department, to my colleagues, friends and family. I am blessed to have crossed paths with you and wish to continue in the future. You have all contributed to this accomplishment.

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TABLE OF CONTENTS

Dedication ...... ii

Acknowledgements ...... iii

List of Tables ...... ix

List of Figures ...... x

List of Abbreviations ...... xii

Preface: Introduction to Dissertation ...... xiv

Chapter 1: Background ...... 1

1.1. Cardiomyocytes ...... 1

1.1.1. Cardiomyocytes in the intact heart ...... 1

1.1.2. Isolated and cultured cardiomyocytes ...... 1

1.2. Cardiac sarcoplasmic reticulum ...... 2

1.2.1. Morphological characterization ...... 3

1.2.2. SR and ER markers ...... 4

1.3. involved in Ca2+ homeostasis ...... 6

1.3.1. Ca2+ homeostasis in the cardiomyocyte ...... 6

1.3.2. Major proteins in the Ca2+ -release complex ...... 7

1.4. Protein: biosynthesis and trafficking ...... 8

1.4.1. Protein synthesis and nascent polypeptide folding ...... 8

1.4.2. Co-translational asparagine (N)-linked glycosylation ...... 9

1.4.3. Post-translational modifications of the N-linked glycans ...... 9

1.4.4. Rough ER and smooth ER ...... 11

1.4.5. ER exit sites and the intermediate compartment ...... 12

1.4.6. Golgi ...... 13

1.4.7. Protein trafficking through the cardiac secretory pathway ...... 14

iv

1.5. Junctional sarcoplasmic reticulum proteins ...... 15

1.5.1. Calsequestrin ...... 15

1.5.1.1. Cardiac muscle calsequestrin (CSQ2) ...... 18

1.5.1.1.1. Ca2+ binding and polymerization ...... 18

1.5.1.1.2. (N)-linked glycosylation ...... 19

1.5.1.1.3. Phosphorylation by protein kinase CK2 ...... 20

1.5.1.1.4. Role of CSQ2 in Ca2+ homeostasis ...... 21

1.5.1.1.5. Characteristics of CSQ2-DsRed ...... 22

1.5.2. Triadin ...... 23

1.5.3. Junctin ...... 23

1.5.4. Junctional sarcoplasmic reticulum protein-protein interactions ...... 24

1.6. Hypertrophy ...... 25

1.6.1. Cardiomyopathies linked to genetic changes in protein components of the Ca2+-release complex in mice ...... 26

1.7. Cytoskeleton ...... 29

1.7.1. Cardiac cytoskeletal components ...... 29

1.7.2. Microtubules and endoplasmic reticulum ...... 29

1.7.2.1. Microtubules ...... 30

1.7.2.1.1. Microtubule disrupting agents ...... 30

1.8. Adenoviral mediated overexpression in cardiomyocytes ...... 31

Chapter 2: Materials and Methods ...... 33

2.1. Animals and heart tissue ...... 33

2.2. Purification of CSQ2 ...... 33

2.3. Cardiomyocyte isolation ...... 34

2.3.1. Surgery ...... 34

2.3.2. Isolation protocol ...... 35

v

2.3.3. Cardiomyocyte culture ...... 36

2.4. Cell treatments ...... 36

2.4.1. Adenovirus infection ...... 36

2.4.2. Nocodazole treatment ...... 37

2.5. Electrophoresis ...... 37

2.5.1. Protein extraction and concentration determination ...... 37

2.5.2. Gel electrophoresis ...... 38

2.5.3. Concanavalin A binding to glycoproteins ...... 38

2.6. Immunological methods ...... 39

2.6.1. Antibodies ...... 39

2.6.2. Immunoblot ...... 39

2.6.3. Immunofluorescence of isolated cardiomyocytes and tissue sections ...... 40

2.6.3.1. Immunocytochemistry on isolated cardiomyocytes ...... 40

2.6.3.2. Immunohistochemistry on tissue sections ...... 40

2.7. Fluorescence microscopy ...... 41

2.8. Quantitative PCR ...... 42

2.9. Mass spectrometry ...... 42

2.10. Statistical analysis ...... 43

Chapter 3: Altered Calsequestrin Glycan Processing is Common to Diverse Models of Canine Heart Failure ...... 44

3.1. Rationale ...... 44

3.2. Results ...... 44

3.2.1. Analysis of CSQ2 glycoforms and phosphoforms ...... 44

3.2.2. Analysis of CSQ2 glycan by ConA binding ...... 49

3.2.3. Steady-state CSQ protein and mRNA levels ...... 50

vi

3.2.4. Immunohistochemical analysis of canine tissue ...... 52

3.3. Summary ...... 54

Chapter 4: Sorting and Trafficking of Junctional SR Transmembrane Protein Across ER Along Microtubules ...... 56

4.1 Rationale ...... 56

4.2 Results...... 56

4.2.1 Species-specific antibodies to junctional SR transmembrane proteins...... 56

4.2.2 Early accumulation of expressed JCTdog and TRDdog at juxtanuclear ...... 58

4.2.3 In situ interaction of JCTdog and TRDdog with CSQ2-DsRed in rough ER ...... 60

4.2.4 Morphological distinctions between smooth ER and junctional SR ...... 64

4.2.5 Inhibition of anterograde trafficking by nocodazole ...... 67

4.3 Summary ...... 69

Chapter 5: Trafficking of the Luminal SR Protein CSQ2 ...... 70

5.1 Rationale ...... 70

5.2 Results...... 70

5.2.1 Detection of CSQ2dog using species-specific antibodies ...... 70

5.2.2 Determination of CSQ2 initial sites of accumulation ...... 71

5.2.3 In situ interaction of TRDdog with CSQ2 ...... 74

5.2.4 Effect of nocodazole on junctional SR sites and CSQ2 trafficking ...... 75

5.3 Summary ...... 76

Chapter 6: Discussion ...... 78

6.1 CSQ2 mass redistribution in two types of cardiomyopathies ...... 78

6.2 Junctional SR subdomains: protein trafficking and accumulation ...... 82

6.3 Model of the inner membrane system in the adult cardiomyocytes ...... 86

vii

6.4 Conclusions ...... 89

Appendix A: Springer License Terms and Conditions ...... 91

Appendix B: Approved Animal Protocol ...... 92

Appendix C: Supplemental Data ...... 93

References ...... 96

Abstract ...... 119

Autobiographical Statement ...... 121

viii

LIST OF TABLES

Table 1-1. CK2 sensitive phosphorylation sites present in the carboxyl- terminal tail of CSQ2 ...... 21

Table 1-2. Summary of junctional SR protein overexpression and suppression studies ...... 27

Table 2-1. Relative reactivity of junctional SR protein antibodies for dog or rat forms ...... 39

Table 3-1. Molecular masses (weighted averages ± S.D.) of CSQ2 from canine heart tissue samples (Da) ...... 46

ix

LIST OF FIGURES

Figure 1-1. Depiction of T-tubules, SR and myofilaments in a mammalian cardiac muscle cell ...... 3

Figure 1-2. Schematic diagram of junctional SR with T-tubule and the Ca2+- induced Ca2+ release process ...... 6

Figure 1-3. N-linked glycan biosynthesis and CSQ2 glycan trimming along the secretory pathway ...... 11

Figure 1-4. Human CSQ amino acid sequences ...... 15

Figure 1-5. Ribbon diagram of CSQ structure ...... 17

Figure 1-6. CSQ polymerization model ...... 19

Figure 1-7. CSQ2-DsRed in adult cardiomyocytes ...... 22

Figure 3-1. CSQ2 masses are altered in heart tissue from mongrel dogs in heart failure ...... 47

Figure 3-2. Further evaluations of CSQ2 mass spectra data ...... 49

Figure 3-3. Con A binding to purified CSQ2 from C and HF tissue samples ...... 50

Figure 3-4. Steady-state levels of CSQ2 protein, mRNA and protein to mRNA ratios in canine hearts ...... 51

Figure 3-5. Detection of CSQ2 in canine tissue slices by immunofluorescence, and CSQ2 antibody reactivity ...... 53

Figure 4-1. Specificity of anti-TRD and anti-JCT antibodies ...... 57

Figure 4-2. JCT accumulation at juxtanuclear sites ...... 58

Figure 4-3. TRD accumulation at juxtanuclear sites ...... 60

Figure 4-4. TRD and JCT move along a common pathway to populate junctional SR ...... 61

Figure 4-5. In situ interactions of JCT and TRD with CSQ2-DsRed at juxtanuclear sites ...... 62

Figure 4-6. CSQ2-DsRed-containing ER traffics between myonuclei in line with MTs ...... 63

Figure 4-7. TRD is retained by its in situ binding to polymerized CSQ2-DsRed in the smooth ER lumen ...... 64

x

Figure 4-8. The CSQ2-binding mutant delTRD readily crosses the smooth ER to concentrate at sites of junctional SR ...... 66

Figure 4-9. Nocodazole disrupts anterograde movement of junctional SR proteins ...... 68

Figure 5-1. Specificity of anti-CSQ2 antibodies ...... 71

Figure 5-2. Time course of CSQ2 accumulation at junctional SR sites ...... 73

Figure 5-3. Higher rates of CSQ2 biosynthesis create higher fluorescence intensities at juxtanuclear sites ...... 74

Figure 5-4. CSQ2 maintained independent transit when co-expressed with TRD and was not similarly disrupted by nocodazole treatment ...... 76

Figure 6-1. Schematic model of junctional SR filling by newly-made junctional SR proteins ...... 88

Figure A-1. Levels of endogenous rat CSQ2 and JCT proteins at various time points in culture ...... 93

del Figure A-2. Time course of TRDdog accumulation at junctional SR sites ...... 93

Figure A-3. The effect of nocodazole concentration and duration of treatment ...... 94

Figure A-4. Junctional SR sites distribution depicted by CSQ2rat endogenous protein localization upon nocodazole treatment ...... 95

xi

LIST OF ABBREVIATIONS

Ad.CSQ2-DsRed: calsequestrin-DsRed adenoviral vector

Ad.CSQ2: calsequestrin adenoviral vector

ANF: atrial natriuretic factor

ATP: adenosine triphosphate

BNP: brain natriuretic peptides

C: control

CAR: coxsackievirus

COP I: cytosolic coat protein I

COP II: cytosolic coat protein II

CPVT: catecholaminergic polymorphic ventricular tachycardia

CSQ: calsequestrin

DAPI: 4’-6-diamidino-2-phenylindole

ECL: enhanced chemiluminesence

ER: endoplasmic reticulum

ERGIC: endoplasmic reticulum-to-golgi intermediate compartment

ESMS: electrospray mass spectrometry

GAPDH: glyceraldehyde 3-phosphate dehydrogenase

GlcNAc: N-acetylglucosamine

HF: heart failure

HFi: Heart failure induced by ischemic microembolisms

HFt: Heart failure induced by tachypacing

JCT: junctin

LV: Left ventricle

xii

MOI: multiplicity of infection

OST: Oligosaccharyl transferase

PBS: phosphate buffer saline

PBS/tween: phosphate buffer saline with 0.2% tween20 pfu: plaque forming unit

RyR: ryanodine receptor

SDS-PAGE: sodium dodecyl sulfate polyacrylamide gel electrophoresis

SERCA: SR/ER Ca2+-ATPase

SR: sarcoplasmic reticulum

T-tubules: transverse tubules

TGN: trans Golgi network

TRD: triadin

xiii

PREFACE

INTRODUCTION TO DISSERTATION

Mechanisms responsible for junctional SR subdomain formation from ER have

remained undefined. This lack of understanding hampers our ability to understand

changes in SR function in disease; such as the dramatic changes in CSQ2 structure

that we validated through a series of measurements made on control and failed heart samples. In this research, we developed a new model of the early cardiac secretory pathway. By defining the individual steps of trafficking for CSQ2 and for its transmembrane binding partners TRD and JCT, we will be able to develop new hypotheses regarding cellular mechanisms of change.

This Dissertation research involves two studies directed at the junctional sarcoplasmic reticulum (SR) and aspects of its dynamic cell biology. In Chapter 3, we describe our investigations of CSQ2 purified from control and failed heart tissue. Mass spectrometric measurements of the intact CSQ2 pool provide detailed data regarding glycosylation and phosphorylation states in vivo. We conclude that the observed changes in cardiomyopathies represent alterations in intracellular trafficking within the

ER of cardiomyocytes. Chapter 4 and 5 describe a series of studies in which we determined the trafficking pathways of newly made junctional SR proteins as a way to better understand the mechanisms underlying changes in protein modification. The studies described in this Dissertation, we believe, provide important new insights into the mechanisms of long-term change in cardiomyocytes that might underlie changes

reported in diseased heart.

xiv 1

CHAPTER 1

BACKGROUND

1.1 Cardiomyocytes

1.1.1 Cardiomyocytes in the intact heart

Cardiomyocytes are specialized cells used to generate force in response to

depolarization of the heart. These cells are large (100-150 m long, 20-35 m wide),

binucleated, and surrounded by a plasma membrane known as the sarcolemma.

Invaginations of the sarcolemma at regular intervals form transverse tubules (T-

tubules), which have roughly a 200 nm diameter. [1, 2]. The T-tubules are not strictly

transverse and can also communicate longitudinally, across several sarcomeres [3].

Intercalated disks spanned by gap junctions join adjacent cells and allow for rapid

conduction of action potentials; hence, all cardiomyocytes contract in synchrony. The

fundamental contractile unit in striated muscle is the sarcomere, defined as the region

confined between two Z-lines. Chains of contractile units comprise myofibrils, and

these contain the contractile thick (myosin), thin filaments (actin and others), and titin

[4]. Contractile force is generated by interaction of actin and myosin, and regulated by

Ca2+ release from intracellular membrane stores.

1.1.2 Isolated and cultured cardiomyocytes

To explore myocardial cell biology, primary cardiomyocytes can be isolated and

cultured. Although neonatal cells are more stable, adult cardiomyocytes are preferred

for cell biological studies because the adult phenotype represents such a unique cell

system [5]. The technique for isolating viable adult ventricular myocytes was first

described in 1976 [6] and adult cells were first cultured in 1977 [7]. Several advantages

to culturing adult primary cardiomyocytes include: ability to carry out longer-term studies

2

that allow manipulation of protein expression [8], examination of a single cardiomyocyte

in a controlled environment, and visualization of their structural components by a series

of well-characterized protein markers. Hence, utilization of cultured adult

cardiomyocytes allows for the extrapolation of results to the in vivo myocardium and

thus, complements intact heart preparations or whole animal studies.

1.2 Cardiac sarcoplasmic reticulum

A network of interconnecting intracellular membranes, the sarcoplasmic reticulum

(SR), is largely equivalent to the endoplasmic reticulum (ER) found in other cell types.

Besides its canonical universal role as the biosynthetic and secretory pathway,

subcompartments of cardiomyocyte ER may surround each myofibril, taking on a similar

pattern in each of the hundreds of sarcomeres in the cell (Figure 1-1). It is this

extensive and unique sarcomeric component of the cardiomyocyte ER that gives rise to

its unique reference as SR. SR is known to play an important role dedicated to storage and release of Ca2+ for regulating muscle contraction and excitation-contraction

coupling. SR is comprised of two distinct functional subdomains, each associated with one of its two major functions: Ca2+ reuptake and Ca2+ release. The site of Ca2+ reuptake remains less well defined morphologically. The primary protein activities associated with these two functions are the SR/ER Ca2+-ATPase (SERCA) and

ryanodine receptor (RyR), but freeze-etch studies suggest that abundant SERCA is

found across sarcomeric membrane structures, including in bilayers close to SR-

sarcolemma junctions [9]. RyR are a second major protein complex, and these have

almost exclusively been described extending as “feet” across the SR-sarcolemma

junctions. The Ca2+ -release complex has been studied using multiple biochemical, biophysical, and microscopic techniques,

3

Since the proteins associated with these two major activities are relatively well

characterized (discussed below), antibodies to these various proteins serve as

important markers for the two subdomains, which has permitted many investigations of

their respective localizations. The vast majority of studies to date have focused on the

Ca2+ release subdomain, and its subcellular morphology. It is clear from many studies

that Ca2+ release occurs from a specialized area of the SR that abuts the T-tubules, and

these sites appear by immunofluorescence as a series of puncta that overlie the Z-disks

of the myofilaments (Figure 1-1).

Figure 1-1. Depiction of T-tubules, sarcoplasmic reticulum (SR) and myofilaments in a mammalian cardiac muscle cell. Each myofibril is surrounded by an intermembrane network of free SR (blue arrow) and junctional SR cisternae (red arrow head) that forms close to T-tubules (green double head arrow), which invaginate the cardiomyocyte near Z- lines (Z).

1.2.1 Morphological characterization

Studies using thin section electron microscopy describe the cardiac intracellular

membrane morphology. Throughout ventricular myocytes, the SR consists of

structurally diverse junctional, corbular and network (also referred to by many as

longitudinal or free) SR. Spanning the region between the T-tubules, a series of

interconnected tubules form the network SR. Specialized junctions of network SR with

sarcolemmal T-tubules form the junctional SR, resulting in close juxtaposition (~15 nm)

of RyR and L-type Ca2+ channels where Ca2+ is released to initiate muscle

contraction[10-12].

A protrusion of the SR network that does not form junctions with the sarcolemma,

4

but expresses RyR, is corbular SR [13, 14]. Corbular SR and non-junctional SR

(prevalent in atria), are not generally thought to be involved in depolarization-induced

Ca2+ release. Non-junctional SR is also the primary source of Ca2+ release in birds [3].

The two major functions of cardiac SR that are typically discussed, Ca2+ release and

Ca2+ reuptake, are therefore not necessarily synonymous with the two

subcompartments of junctional and free SR. The various terminologies reported to date

are descriptive of the overall sarcomeric morphology, but more detailed morphological

descriptions may not be warranted until a better understanding of the relationship

between SR and the biosynthetic/secretory pathway can be established. Even less clear is how junctional SR and free SR subcompartments are maintained as dynamic

subcompartments of the cardiomyocyte secretory pathway.

1.2.2 SR and ER markers

The ER is a multifunctional organelle in all cells, and the vast majority of proteins

are ubiquitously present in all cell types. In cell types other than muscle, ER is probably

considered more for its roles in protein biosynthesis and secretion. Among the more

abundant proteins found in all cells are a group of luminal resident proteins that can be

generally referred to as "reticuloplasmins” [15, 16]. Whereas luminal proteins should be

secreted through bulk flow, the reticuloplasmins are actually retained in the ER. Over

time, it has became understood that most, if not all, of these reticuloplasmins are

involved in the folding of other proteins inside the secretory pathway. This function has

become generally known as "chaperone" activity. The prototypical member of the

family is GRP78 (glucose-regulated protein, MW 78 kDa), also commonly known as the

protein BiP (Binding-immunoglobulin protein), named for its role in assisting folding of

antibody molecules in the B-cell ER [17]. GRP78 may have a wide role in folding

5

nascent polypeptides as they are translated, serving as a template upon which proteins

fold in conjunction with the aqueous environment of the ER lumen. With the cloning of

several of these ER resident proteins, it was discovered that a common C-terminus

tetrapeptide (-KDEL) was responsible for its ER retention, and a KDEL receptor(s) is

now well characterized that retrieves these chaperones back to ER [18]. These findings

helped lead to the understanding of basic cell biology, as it was discovered that a

retrograde trafficking pathway was present in all cells (see Section 1.4.1).

Myocyte ER/SR contains both the common ER reticuloplasmins (Cala and Jones

1994) along with similar myocyte specific proteins, calsequestrin (CSQ), sarcaluminin,

and histidine-enriched Ca2+-binding protein [16]. CSQ alone is enriched and localized to

the junctional SR subcompartment. None of the myocyte enriched luminal ER/SR

proteins, however, contain the KDEL retrieval sequence, and so the mechanisms for

their ER/SR enrichment have not been understood. In the case of CSQ, data suggests

that it is retained in particular subcompartments due to its unique and subcompartment-

dependent Ca2+-dependent polymerization [19]. Comparable mechanisms may be

responsible for the concentration of sarcaluminin and histidine-enriched Ca2+-binding protein in free SR. Because of the abundance of the reticuloplasmins in the ER and

SR, they are often used as protein markers of ER and SR.

The locations within the cardiac ER in which protein biosynthesis occurs, and areas from which SR proteins are trafficked to ER subdomains (e.g. free versus junctional SR) have remained largely undefined. Studies have shown that diffusion of

Ca2+ occurs between ER and SR compartments suggesting that as a Ca2+-handling system, there exists a breadth of functional domains across the biosynthetic/secretory pathway [20].

6

1.3 Proteins involved in Ca2+ homeostasis

1.3.1 Ca2+ homeostasis in the cardiomyocyte

2+ 2+ Rises in the levels of intracellular Ca in the cytoplasm ([Ca ] in) during systole are highly regulated, and regulation of Ca2+ cycling is fundamental for achieving maximal contraction. When plasmalemma and T-tubules are depolarized, a Ca2+ current flows through the L-type Ca2+ channels. The Ca2+ that enters the cell through

the L-type Ca2+ channels (trigger Ca2+) as a result of depolarization, binds to the RyR, and triggers the release of a larger flux of Ca2+ (activator Ca2+) from the SR into the

cytoplasm. This process is termed Ca2+ -induced Ca2+ release [4, 21] (Figure1-2). The magnitude of Ca2+ release depends on the rate of filling of the SR with Ca2+, the time

interval between successive release events, and the magnitude of the trigger Ca2+ [22].

The rise and fall of Ca2+ that precedes every contraction is known as the Ca2+ transient.

Figure 1-2. Schematic diagram of junctional SR with T-tubule and the Ca2+-induced Ca2+ release process. The protein transmembrane components of junctional SR: triadin (TRD), junctin (JCT) and luminal calsequestrin (CSQ) bound to Ca2+ (red spheres) associated with the Ca2+ -release channel (ryanodine receptor, RyR2). Upon depolarization Ca2+ enters through the L-type Ca2+ channel, triggers the release Ca2+ from the SR through RyR, a process known as Ca2+ induced Ca2+ release, leading to contraction. During relaxation, SR/ER Ca2+ ATPase (SERCA) reuptakes Ca2+ to restore the Ca2+ levels in SR (Modified from [26].

2+ 2+ Ca transients lead to Ca binding to troponin C (Km~0.1 μM) [23] inducing

subsequent conformational changes to activate actinomyosin ATPase, which result in

development of force. With activation of striated muscles, the troponin-tropomyosin

complex on the thin actin filaments transduces chemical energy into mechanical force

7

[24] and exposes myosin binding sites, thus, allowing formation of cross bridges, conversion of ATP chemical energy into force, sliding of the filaments, and shortening of the sarcomeres [25].

The released Ca2+ is removed from the myofilaments and cytosol, into the SR

lumen by SERCA, into the extracellular space by plasma membrane Na+,Ca2+-

exchanger and plasma membrane Ca2+-ATPase, or into the mitochondria via a Ca2+ uniporter. Over the course of many depolarizations, in order for the cell to remain in a steady state with respect to cellular Ca2+ balance, the efflux and influx of Ca2+ must remain equal [4].

1.3.2 Major proteins of the Ca2+ -release complex

A quaternary complex of four proteins: RyR, triadin (TRD), junctin (JCT) and

CSQ, is thought to be primarily involved in the coordinated operation of Ca2+ release,

which has been proposed based on immunoprecipitation and fusion-protein pull down

assays [27]. Physical coupling of CSQ, TRD and JCT modify the resting and voltage-

activated Ca2+ release through RyR, based on junctional SR Ca2+ load, suggesting that these proteins play an active role in relaying information to RyR about the Ca2+ load in the SR [28]. Other studies, including studies of TRD and JCT knockout mice, have suggested that these proteins may play a more structural role. The known problem with studies of transgenic and knockout mice has been the coordinated changes in other junctional SR proteins (section 1.6.1)

RyRs mediate the release of Ca2+ from the SR, which is an essential step in

cardiac muscle contraction [29]. The RyR is a homotetramer, with a predicted size of

564 kDa in humans, encoded in mammals by three with strong sequence

homology between the isoforms: RYR1 (first to be sequenced and cloned in 1989) [30]

8

is expressed in skeletal muscle, RYR2 in cardiac muscle [31], and RYR3 in smooth

muscle and the brain [32]. Between the T-tubule and terminal cisterna membrane, the

cytoplasmic domain of RyR2 forms a major part of the junctional complex [33].

In cardiac muscle, mutations in the RYR2 are associated with the dominant genetic form of catecholaminergic polymorphic ventricular tachycardia (CPVT), a familial arrythmogenic disease that manifests in childhood and adolescence [34] and is characterized by syncope and sudden death following exercise and emotional stress.

RYR2 mutations result in “leaky” RyR2 channels by yet unknown mechanisms. The

inappropriate Ca2+ release results in increased Na2+, Ca2+ exchanger activity, which

depolarizes the cell membrane, giving rise to delayed after depolarizations [35].

Mutations in the RYR2 gene are also associated with stress-induced polymorphic ventricular tachycardia and arrhythmogenic right ventricular dysplasia [36].

1.4 Protein: biosynthesis and trafficking

1.4.1 Protein synthesis and nascent polypeptide folding

Protein translation from mRNA is carried out on ribosomes. Large (60S) and small (40S) ribosomal subunits assemble at the AUG codon of mRNAs at the 5’-end and disassemble at the UAG/UGA/UAA stop codons at the 3’-end. Cytosolic, nuclear and peroxisomal proteins are synthesized on cytosolic ribosomes [37]. However, during translation, short hydrophobic peptide signal sequences direct a subset of nascent polypeptides to assemble on ribosomes that are bound to the ER. Proteins translated on rough ER include those destined for the secretory and endocytic compartments (ER,

Golgi, endosomes and lysosomes), for the plasma membrane and for secretion to the extracellular space. The signal sequences for these cellular proteins bind the signal recognition particle in the cytoplasm, thus slowing down chain elongation [38, 39] until

9 the ribosome engages a proteinacous channel, the translocon, formed by two Sec61αβγ heterotrimeric complexes at the ER membrane [40, 41]. With the formation of a translocon, the protein synthesis is resumed, the short hydrophobic signal sequence may then be removed (for some proteins such as CSQ2), and nascent chains co- translationally enter the ER lumen [42].

1.4.2 Co-translational asparagine (N)-linked glycosylation

The most prominent modification of secretory proteins is N-linked glycosylation, which occurs for a very high percentage of membrane and secreted proteins [43].

Oligosaccharyl transferase (OST) transfers preassembled glycans, composed of three glucoses, nine mannoses, and two N-acetyl glucosamine residues (GlcNAc2Man9Glc3), from a lipid pyrophosphate donor in the ER membrane, dolichol-PP, to a nascent polypeptide chain [44]. The transfer to the consensus sequence Asn-X-Ser/Thr generally occurs co-translationally, once it has emerged 12-14 amino acids (30-40 Å), aligning the Asn with the active site of OST in the ER lumen [45]. An inhibitor of N- linked glycosylation, tunicamycin, leads to improper or incomplete folding, and failure to reach native conformation for proteins that do not pass ER quality control [46]. These proteins are retained in the ER [47], and eventually degraded [48], a process termed ER associated degradation. A mature properly folded protein contains the glycan

GlcNAc2Man9, but this glycan will be reliably and predictably trimmed and modified as it traverses the secretory pathway.

1.4.3 Post-translational modification of the (N)-linked glycans

Early quality control mechanisms are involved in the trimming of glucose from ER glycoproteins. The enzyme glucosidase I removes the first glucose of the glycan core, followed by the removal of the second glucose by glucosidase II. Molecular chaperones

10 known also as mammalian lectins (such as calnexin and calreticulin, discussed Section

1.4.1) can bind to the monoglucosylated core of the glycoprotein, exposing it to a thiol- disulfide oxidoreductase, ERp57, for proper disulfide bond formation during the on going folding process. Glucosidase II removes the remaining glucose residues to release the bound chains from calnexin and calreticulin. Incompletely folded proteins get reglucosylated by UDP-Glc: glycoprotein glucosyltransferase, which serves as a folding sensor until the glycoprotein is properly folded or degraded [49].

Removal of mannose residues occurs first by ER α1,2-mannosidase I [50, 51], a class I mannosidase [52], which catalyzes the first mannose trimming step concurrently or immediately following glucose trimming. ER α1,2-mannosidase I yield a specific

Man8GlcNAc2 isomer from Man9GlcNAc2, by cleaving a single α1,2-mannose residue.

The enzyme is inhibited by kifunesine and 1-deoxymannojirimycin [52]. In rat hepatocytes and several other mammalian cell lines, an additional mannosidase resistant to kifunesine, termed ER α1,2-mannosidase II, has been identified and shown to produce a Man8GlcNAc2 isomer, distinct from that of ER α1,2-mannosidase I [51,

53].

It is known that CSQ2 undergoes extensive mannose trimming from Glc2Man9

(Man9) down to Man3-4 [54, 55]. However, these authors have argued that post-ER glycan trimming of CSQ2 would be expected to add terminal GlcNAc to the CSQ2 protein (Figure 1-3). The fact that CSQ2 molecules do not contain any additional sugars, but instead only undergoes mannose trimming is consistent with the idea that

CSQ2 molecules never leave the ER, and that only ER α-1,2 mannosidase is expected to act upon CSQ2. Characterization of mannosidases in individual cell types has not been carried out, and little data exist on the distribution of mannosidases in myocytes,

11 and so the connections between CSQ2 glycan structure and its intracellular trafficking

remain uncertain.

Figure 1-3. N-linked glycan biosynthesis and CSQ2 glycan trimming along the secretory pathway. From right to left: glycan synthesis begins at the cytoplasmic side of the ER membrane and is then flipped to the luminal side of the membrane. Once the formation of the core oligosaccharide (Glc3Man9GlcNAc2) is complete, the enzyme OST (not shown) catalyzes the glycosylation of the Asn residue in the nascent growing peptide. When the glycoprotein has properly folded (grey oval) the process of mannose trimming occurs by intracellular mannosidases as the protein traffics through the ER and Golgi. The glycan of CSQ2 is highly mannose trimmed contains only a variable number of terminal Man residues, and no additional sugars are added, indicating that it does not reach the Golgi, but is retained within junctional SR to perform its physiological function. (Modified from [43]

1.4.4 Rough ER and smooth ER

The ER is a membrane protein biosynthesis and maturation compartment topologically equivalent to the extracellular milieu [56]. It is classically subdivided into rough ER areas that are sheet-like and ribosome studded and smooth ER that is more tubular in structure. ER domains are continuous, forming an intricate network continuously undergoing rearrangements, composed of fenestrated cisternae and

anastomosing tubules through most of the cytoplasm [57].

Rough ER a highly dynamic part of the ER organelle marked by the site of

protein assembly and includes the translocon, a large complex of proteins that make up

the entry point into the secretory pathway. As discussed above (section 1.2.2), high

12 concentrations of molecular chaperones in the ER are thought to assist protein maturation and prevent unfolded chain aggregation. Rough ER contains homologs of classical chaperone members of the Hsp70 and Hsp90 family, namely GRP78/BiP [58] and GRP94 [59]. There are many other chaperones including lectin-like molecular chaperones, such as calnexin, calreticulin and calmegin [60], which recognize the core oligosaccharide on nascent proteins, and further assist in proper folding of ER glycoproteins. Resident ER luminal proteins discussed above, such as GRP78/BiP,

GRP94 and protein disulfide isomerase, which covalently modifies newly synthesized proteins, share a common carboxyl-terminal tetrapeptide sequence, the KDEL motif, which is necessary for retention of luminal ER proteins and sufficient to cause a normally secreted protein to be retained [61]. Retrograde transport of KDEL containing proteins, from the intermediate compartment and the cis Golgi, is mediated by a membrane-bound KDEL receptor [62].

1.4.5 ER exit sites and the intermediate compartment

A great deal of research over the past 10-20 years has established a basic mechanism by which folded proteins exit the ER to continue across more distal compartments of the secretory pathway. Transitional ER or ER exit sites form at small ribosome free regions within the rough ER from which cytosolic coat protein II (COPII) vesicles arise [63, 64]. COPII vesicles package properly folded and assembled proteins and transport them out of the ER into the Golgi-directed pathways. A COPII coated bud that transport cargo anterogradely, is initially formed by Sar1p recruited to the ER

membrane followed by Sec23-Sec24p and finally Sec12-Sec31p [65]. These early

proteins of the ER exit apparatus, which are recruited to ER exit sites, are often used as

markers. Sec23 has been used as a marker for ER exit sites in cultured

13 cardiomyocytes [66]. In the adult cardiomyocyte, it remains to be established where ER exit sites are found, and whether SR formation from ER might require ER exit.

These COPII transport vesicles fuse to generate an ER-Golgi intermediate compartment (ERGIC) in mammalian cells. While the ERGIC may consist of discrete intracellular structures, it has little functional interest, other than its role as a step that is both post-ER and pre-Golgi. Once in Golgi, proteins are transported between

compartments via COPI coated vesicles [67].

1.4.6 Golgi

COPII vesicles exit the ER to transfer lipid and membrane protein to cis Golgi cisternae. An important and established activity of the Golgi is the modification of glycan moieties, which also serves as a marker of the transfer of a glycoprotein from the ER to the Golgi and beyond. Numerous studies over time have focused on the processing of

N-linked glycoproteins continues in the Golgi. In addition, glycan modifying enzymes such as glycosyltransferases, glycosidases, and nucleotide sugar transporters are present in a defined manner within Golgi compartments such that each acts consecutively on specific substrate(s) generated earlier in the pathway.

GlcNAc moieties are commonly added to free hydroxyl termini of N-linked glycans by the enzyme GlcNAc transferase. While this modification is likely to involve widespread changes in the functions of individual glycoproteins, its determination in individual proteins is often used by scientists to establish basic trafficking steps.

Terminal GlcNAc moieties are thought to promote resistance to the enzyme endoglycosidase H, compared to high mannose oligosaccharides added in the ER.

Cisternal migration moves proteins destined to be secreted to the trans face of the Golgi and subsequently into a complex network of vesicles, termed trans Golgi network

14

(TGN). The TGN contains multiple sorting domains and produces clathrin-coated

vesicles, but not COPI vesicles, whereas earlier Golgi cisternae produce COPI vesicles.

In addition to serving as the exit face of the Golgi, the TGN is the interface between the

Golgi and the endocytic system. Clathrin cooperates with adaptors to produce vesicles

that carry certain cargoes from the TGN to endosomes [68]. Secretory cargoes are

delivered from the TGN to the plasma membrane by a variety of routes. Therefore,

although protein N-linked glycosylation and other post-translational modifications in the

various organelles are of fundamental importance in protein function in the cell, the

experience gained has established glycoproteins as reliable measures of protein

trafficking

1.4.7 Protein trafficking through the cardiac secretory pathway

For proteins to exert their functions, proper subcellular localization is essential.

Human diseases have been linked to aberrant localization of proteins caused by

dysregulation of protein trafficking [69]. Basic anterograde flow through the secretory pathway involves proteins within membrane vesicles moving along default cytoskeletal pathways, either to be secreted (the case for luminal proteins) or to plasma membrane

(for transmembrane proteins).

Yet, the intracellular membrane trafficking pathways and targeting mechanisms involved in the final localization of junctional SR proteins remain unknown. One reason for this is the fact that cardiomyocytes are not the subject of wide study of cell biologists due to their unique structure. Meanwhile, cardiac SR biologists have not investigated protein trafficking due in large part to the enormous effort and expertise of laboratories in studying the details of the Ca2+ release event, at the cost of ignoring the longer-term

changes that would occur with changes in cell biology. Given that cardiac pathologies,

15 and loss of contractile strength in the myocardium, are likely to involve more chronic cellular changes, these deficits in understanding form critical needs for research.

1.5 Junctional sarcoplasmic reticulum proteins

1.5.1 Calsequestrin

CSQ is an abundant myocyte protein, and major Ca2+-binding protein in the SR

lumen of cardiac and skeletal muscle. The fast-twitch skeletal muscle isoform (CSQ1)

was first discovered in 1971 [70, 71] and is encoded by CASQ1 gene [72]. The protein

in heart, encoded by CASQ2 gene [73], was not discovered until 1983. The two

sequences exhibit 65% identity (Figure 1-4), and both are highly acidic proteins

(isoelectric point, pI ~ 4), due to more than 37% Asp and Glu residues in the sequence.

Figure 1-4. Human CSQ amino acid sequences. Sequences of human cardiac (black) and skeletal (red) muscle CSQ isoforms are 65% identical. The site of N-linked glycosylation (purple shaded box) is conserved in both isoforms. A specific cardiac isoform tail (cyan shaded box) contains two CK2-sensitive serine (S) phosphorylation sites in humans (NCBI database). CSQ1, like CSQ2, undergoes glycosylation on 316Asn. Because it does not contain the unique C-terminus of CSQ2, it is not phosphorylated in vivo. It can be phosphorylated in vitro by protein kinase CK2 on Thr373, a site that is not phosphorylated in vivo [74].

16

CSQ1 was originally discovered as a Ca2+ -binding protein component of muscle

microsomes, localized to the lumen of the skeletal muscle SR membrane. The mass of

the protein is similar for CSQ1 and CSQ2, having an approximate molecular weight of

42,000 Da. Both CSQ1 and CSQ2 migrate aberrantly on SDS-gels (Laemmli reagents),

with CSQ1 migrating as if a 63 kDa protein, while CSQ2 runs at 55 kDa. These aberrant mobilities, related to the low SDS-binding of the protein, contributed to the idea

prior to 1983 that CSQ did not exist in the heart. When CSQ1 and CSQ2 are compared

form the same species, CSQ1 is always slightly smaller than CSQ2 because it lacks a

20-25 amino acid C-terminus. CSQ1 is exclusively found in fast-twitch (type II) muscle

fibers, while CSQ2 exists in slow-twitch fibers [75].

The three dimensional structure of CSQ was first reported by Wang et al

(Figure1-5), showing that the monomeric form of CSQ is made of three domains; I

(residues 12-124), II (residues 125-228) and III (residues 229-352), which are nearly

highly similar to the functional topology of bacterial thioredoxin, however, CSQ has not

been shown to exhibit thioredoxin activity itself. Each CSQ domain consists of four α-

helices that line (two on each side) a hydrophobic core, consisting of five β-sheets. The

domains are connected by short sequences, which along with the secondary structural

elements (α-helices), consist of acidic residues that mark the overall protein as

hydrophilic [76].

Kang and co-workers [77] have also compared the surface charge densities for

CSQ1 and CSQ2 and shown clear differences that may account for differences in their

polymerization characteristics (section 1.5.1.1.1). These differences, however, cannot

be immediately translated into differences in their biochemistry and biology because

CSQ-interaction proteins (for example, TRD and RyR) are also generally tissue specific,

17 as is the structure of the sarcomeric SR compartments.

Figure 1-5. Ribbon diagram of CSQ structure. Monomeric CSQ2 consists of three domains, with the phosphate molecules (red triangle) schematically attached to serine residues 378, 382 and 386 in the (canine) cardiac-specific tail (red rectangle). The glycan is also schematized: 9 mannoses (blue circles) and 2 GlcNAc (purple squares) is attached to Asn316 (Modified from [76]).

1.5.1.1 Cardiac muscle calsequestrin (CSQ2)

CSQ2 is a major protein constituent of junctional SR in hearts that functions in

Ca2+ release through its effects on the RyR2 via the two transmembrane proteins TRD

and JCT [78-80]. In spite of decades of research, the functional role of CSQ2 in the

heart remains unknown. It was long thought to act as a Ca2+ buffer based upon its ability to bind very large amounts of Ca2+ in vitro (30-50 mol Ca2+/mol CSQ), however, studies of CSQ2 depletion in mice showed that levels of SR-releasable Ca2+ were not altered. Studies described in this dissertation are not meant to determine the function of

CSQ2, however, elucidation of novel functions of its cell biology, in combination with an examination of its changes in structure in animals with heart failure, offer interesting new insights into its possible role in maintaining structural features of the cardiomyocyte.

1.5.1.1.1 Ca2+ binding and polymerization

CSQ2 undertakes major changes in structure upon Ca2+ binding, and converts from a “sticky” conformation, which binds stably to phenylagarose in micromolar Ca2+, to

18 a hydrophilic conformation in millimolar Ca2+, which causes it to immediately unlatch

from phenylagarose (Cala and Jones, 1983). This Ca2+-dependent conformational change is widely used to purify CSQ2 from crude extracts (Cala and Jones, 1983;

Jacob et al, 2012). Cloning of CASQ revealed that the encoded protein does not contain a distinct Ca2+ binding domain as part of its secondary structure. From the

crystal structure, it was determined that Ca2+ binds to CSQ2 through electrostatic

attractions to negatively charged residues that are, exposed on the surfaces. At low

Ca2+ concentration CSQ2 is present as a soluble monomer. As the Ca2+ concentration

increases, front-to-front surface interactions occur, followed by back-to-back surface

interactions (Figure 1-6) as more Ca2+ is required to shield the negatively charged

residues present at the C-terminus [77].

The segregation of CSQ2 to junctional SR has been hypothetically accounted for

by TRD and JCT transmembrane proteins restricted to junctional SR [27, 81-83].

However, overexpression studies of CSQ in non-muscle cells show clearly that

polymerization can determine localization [19, 84]. CSQ2 is very efficiently retained

within the ER lumen of all non-muscle cells examined to date. Its strict localization is

confirmed by its glycan structure, which contains only Man9 and Man8 glycans.

Insertion of epitope-tags was found to reduce ER retention compared to wild-type

CSQ2, suggesting that these disrupted forms have undergone a loss of the sequence-

specific polymerization that is necessary for proper localization [84]. Finally, CSQ1

does not concentrate in the ER of non-muscle cells, instead CSQ1 efficiently

polymerizes along microtubules (MTs) that emanate from the microtubule-organizing

center. CSQ1 glycans undergo a more substantial mannose trimming (Man 5-9). Thus,

CSQ2 and CSQ1 each polymerize, but they concentrate in either ER or ERGIC,

19 respectively [19]. It is hypothesized that in normal heart, CSQ2 is also retained in junctional SR by this subcompartment-specific polymerization [85].

Figure 1-6. CSQ polymerization model. A representation of face-to- face (through N-terminal arm) interactions of CSQ monomers, followed by back-to-back interactions (through C-terminal tail), as Ca2+ concentration increases. CSQ eventually forms a linear polymer with the front-to-front and back-to-back interactions (Modified from [86]

1.5.1.1.2 N-linked glycosylation

N-linked glycosylation (section 1.4.2) of CSQ2 and CSQ1 occurs on Asn316

(Figure1-3 and 1-4), which is a conserved residue among species and glycoforms [87].

In non-muscle cells, CSQ2 exists in two glycoforms (Man8- and Man9-GlcNAc2); however, CSQ2 purified from tissue (native) exists in a larger range of glycoforms

(Man1-4GlcNAc2), exhibiting a high degree of mannose trimming compared to most proteins [49, 54], suggesting that CSQ2 remains inside the classical mammalian ER compartments, where mannosidases are thought to be involved in glycan trimming

(section 1.4.3). It is therefore expected that CSQ2 does not traffic to early Golgi compartments, because GlcNAc residues would otherwise be expected to be present on CSQ2 glycans.

The discovery that CSQ2 glycan structure is reproducibly altered in heart tissue from beagles induced into failure by tachypacing [55] suggested that CSQ2 trafficking was altered. The CSQ2 glycan present in failed hearts are comprised of two general structures: those with relatively standard, highly trimmed structures (Man4-7), and a new population of molecules not present in normal dog heart tissue. These new glycan

20 structures show a lack of mannose trimming (Man9,8) expected for newly made proteins in the ER (discussed in Chapter 3).

1.5.1.1.3 Phosphorylation by protein kinase CK2

The carboxyl terminus of canine CSQ2 molecules extends an additional 20-25 amino acids from the corresponding CSQ1 sequence (Figure 1-4). This cardiac-specific tail is highly phosphorylated in vivo by protein kinase CK2 (formerly casein kinase II), on a cluster of three serine residues (Ser 378,382,386) in the canine form of CSQ2, but two

serine residues in other species (Table 1-1) [66, 74]. Consisting of two tightly

connected β subunits and two α/α’ subunits, protein kinase CK2 is present in all eukaryotic cells [88].

The presence of phosphate on purified CSQ2 has been shown by multiple experimental approaches [54, 74], but the clearest indication is the presence of mass

isoforms that differ only in phosphate content. Following removal of the CSQ2 glycan

from purified CSQ2 using endoglycosidase H, mass spectrometry of intact CSQ2

protein from heart tissue is composed of only four mass peaks, corresponding to 0, 1, 2,

or 3 phosphates, each differing by about 81 Da. The stoichiometry of canine CSQ2

phosphorylation is ~1.5 mol Pi/mol CSQ, but the phosphates are not evenly distributed

among CSQ2 molecules. CSQ2 is thought to undergo constitutive phosphorylation in

the early secretory pathway compartments and dephosphorylation in distal ones [89].

Phosphorylation has been hypothesized to occur in the rough ER, due to the fact that higher phosphorylation levels were observed in the glycoforms characteristic of proximal secretory compartments [89].

In contrast to more commonly studied hormone-mediated phosphorylation reactions, CSQ2 phosphorylation by CK2 has been hypothesized to result from a

21 process that occurs co-translationally. The data that supports this hypothesis is as follows: 1) CK2 acts as the CSQ2 kinase in intact cells, but can theoretically only co-

localize with CSQ2 prior to translocation across the ER during its biosynthesis [66], 2)

CSQ2 phosphorylation is maximal for CSQ2 molecules with unprocessed glycans, and

decreases in relation to extent of mannosidase processing [89] and 3) phosphorylation

in situ is only measurable when CSQ2 is highly expressed under cytomegalovirus

(CMV) promoter, consistent with the requirement for a co-translational reaction [89].

This phosphorylation reaction has been validated by another lab [90], but has primarily

been studied by the Cala laboratory.

Table 1-1. CK2 sensitive phosphorylation-sites present in the carboxyl-terminal tail of CSQ2.

Sequences from the C-terminal cardiac specific tail are listed, with serine residues (S) as the sites phosphorylated by CK2 for different species. For rat & dog.

1.5.1.1.4 Role of CSQ2 in Ca2+ homeostasis

CSQ2 is the most abundant Ca2+ binding protein in cardiac membrane, although

several Ca2+-binding proteins such as histidine-rich Ca2+ binding proteins and

sarcaluminin are also present in high levels in SR lumens [16]. From the time of its

discovery, CSQ was predicted to act as a Ca2+ buffer and storage protein [91]. Earlier

acute overexpression studies in cultured adult rat cardiomyocytes have supported this

hypothesis, since doubling the levels of CSQ2 in cultured cells produced enhanced Ca2+ transients [92], and increased SR Ca2+ content (load), determined as 10 mM caffeine

22 releasable Ca2+. This was the first study to offer physiological data that the Ca2+ released during muscle contraction emanates from CSQ2, as the luminal Ca2+ storage protein. Studies of CSQ2 in bilayer experiments have shown that CSQ2 led to increased RyR2 open probability [28, 93]. Such data suggested that, independent of any effects on SR Ca2+ buffering, CSQ2 could act directly on RyR2, and that Ca2+ release may be regulated by CSQ2.

1.5.1.1.5 Characteristics of CSQ2-DsRed

DsRed, a red fluorescent protein cloned from the Discosoma coral [94], is an obligate tetramer in vitro [95, 96] and in living cells [96], confirmed by a variety of techniques. A unique subcellular localization of the fluorescent fusion protein CSQ2-

DsRed [19] around the nucleus was observed following its overexpression in cultured adult rat cardiomyocytes. The ability to distinguish the red fluorescent CSQ2 polymer from the more mobile monomer was made possible by the immunoreactivity of monomers, and loss of immunoreactivity in the CSQ2-DsRed polymer. The resultant pattern showed that the CSQ2-DsRed polymer first accumulates in perinuclear rough

ER, and is only able to traffic anterogradely as a monomer (Figure 1-7).

Figure 1-7. CSQ2-DsRed in adult cardiomyocytes. When CSQ2 is fused to the fluorescent protein DsRed it forms tetramers that fluoresce red. These tetrameric CSQ2-DsRed complexes are, interestingly, non-immunoreactive with either anti-CSQ or anti-DsRed (green). (A) Within 20 h overexpression of CSQ-DsRed adenovirus, anti-DsRed detects CSQ fusion protein around the nucleus that slowly traffics peripherally. (B) By 48 h in culture, DsRed tetramers are fluorescing around the nucleus as protein levels build up. Meanwhile, the monomeric CSQ2-DsRed continues trafficking peripherally (Modified from [85].

23

1.5.2 Triadin

TRD is a transmembrane protein of junctional SR that binds to CSQ and may anchor CSQ to the RyR. It was first identified in 1991 as a 95 kDa integral membrane protein enriched in junctional SR vesicles from skeletal muscle, but a smaller, alternatively spliced form was reported in heart in 1996 [97]. The majority of the protein sequence (C-terminus) is localized in the SR lumen, with a single membrane spanning domain and a short N-terminal positively-charged cytoplasmic segment [81, 98] that binds on one side of a “KEKE” motif within a beta sheet structure [99].

The primary isoform expressed in cardiac muscle is triadin-1 (TRD in this manuscript), a shorter splice variant of skeletal muscle triadin, arising as an alternative splicing from a common gene. For the first 257 amino acids, triadin in cardiac and skeletal muscle are identical. However, cardiac muscle TRD incorporates a unique 21 amino acid cardiac specific tail [97, 99]. On SDS-PAGE, canine cardiac TRD appears as a doublet, the glycosylated and unglycosylated forms resolving at 40 kDa and 35 kDa, respectively [99]. Mouse cardiac TRD also occurs in both glycosylated and unglycosylated forms [100]. It has been shown in a non-muscle cell model that the unglycosylated form of triadin is more rapidly being broken down by the proteasome

[101]. The glycosylation site (75Asn) is only 7 residues from the ER membrane surface,

which makes its glycosylation very inefficient. Why the TRD sequence has maintained

this inefficient level of glycosylation through its evolution remains an enigmatic feature

of its structure and cellular processing.

1.5.3 Junctin

In 1995, JCT was purified and its primary structure determined; it exhibits

significant sequence identity to the liver ER aspartyl β-hydroxylase, representing a

24 unique splice variant present in striated muscle cells. It has limited homology with rabbit skeletal muscle TRD, especially in the transmembrane region (type II signal anchor domain). JCT is enriched in junctional SR membranes from skeletal and cardiac muscle [102]. JCT consists of 210 amino acids with a molecular mass of 26 kDa; it has a short amino acid cytoplasmic N-terminus, a single transmembrane domain and a highly charged C-terminal luminal tail that can bind CSQ2 [103]. The protein in heart does not from disulphide-linked oligomers and does not appear to be glycosylated [103].

1.5.4 Junctional sarcoplasmic reticulum protein-protein interactions

TRD has been shown to bind CSQ [104], RyR [81] and interact with JCT [27] through a common luminal domain. The binding sites for CSQ [104] and RyR [105] contain a KEKE motif localized within residues 210-224. Since electron microscopy of intact muscle shows CSQ adherent to the luminal face of the RyR [106] and TRD has the ability to bind both RyR and CSQ, it can be inferred an anchoring function for TRD of CSQ.

Immunofluorescence and immunoblotting show that JCT co-localizes with CSQ and RyR [103]. In junctional SR vesicles, JCT has been shown to be the major CSQ binding protein in SR [102]. The high levels of basic amino acids were predicted to be important for JCT’s ability to bind to CSQ to provide a membrane anchor for CSQ [73].

The CSQ binding domain on JCT was localized by fusion protein analysis to the intraluminal region amino acid residues 46-210, the region containing the charged amino acids [27]. In the same study, JCT was shown to interact with CSQ, and the RyR in a Ca2+ dependent manner and with TRD in a Ca2+ independent manner. Other

studies have more recently suggested that JCT and TRD are not simply redundant, but

might have distinct properties and detailed functions in the myocyte [83].

25

1.6 Hypertrophy

When heart muscle get progressively weaker and can no longer pump enough blood to meet the metabolic needs of the body, heart failure (HF) can develop from such

ventricular dysfunction. Heart failure may also result from one or the sum of many

causes. Such a progressive disorder must be managed in regard to not only the state

of the heart, but to the condition of the circulation, lungs, neuroendocrine system and

other organs as well.

Associated with ventricular dysfunction are alterations in cardiac size, shape and

function. These changes can result from changes in specific genomic expression,

molecular mechanisms, or subcellular structures. Recent studies have observed that T- tubule integrity is lost in a mouse model of ventricular hypertrophy and HF [107]. The reasons for T-tubule reorganization are under active investigation. One current focus of study is the junctional SR protein junctophilin-2, a transmembrane protein that is able to interact with phosphoinositides in the T-tubules.

As a direct activator of myofilaments and thus contraction, Ca2+ is fundamental

for heart function. In the failing heart, there can exist an inability to properly regulate the

Ca2+ transient or Ca2+ interaction with contractile proteins. Changes in the steady-state

distribution of Ca2+ within the cardiomyocyte of stressed and failing hearts are thought

to underlie reduced contractility [108-110]. Candidate mechanisms for reduced

contractility include changes that prevent normal filling of SR Ca2+ stores, normal

release of Ca2+ [109, 110], and Ca2+-dependent changes in transcription [111-115].

We recently showed that CSQ2-DsRed protein accumulation at juxtanuclear sites could strongly stimulate local spontaneous release of Ca2+ through RyR [116, 117].

This compartment is also the putative site at which CSQ2 is co-translationally

26 phosphorylated by protein kinase CK2 [66]. Concentration of CSQ2 in this juxtanuclear

subcompartment might contribute to a functional Ca2+ microdomain that supports a

cardiac remodelling phenotype [108, 113].

1.6.1 Cardiomyopathies linked to genetic changes in protein components of the

Ca2+-release complex in mice

Mouse models of the protein components of the Ca2+-release complex, while

validating possible roles of CSQ2 in Ca2+ release, have not been successful in resolving

molecular mechanisms of Ca2+ release or junctional SR formation. Beyond the

complexities that underlie the likely molecular mechanisms, mouse genetic models have

ended up leading to changes in multiple protein components (Table 1-2).

Mechanical stress, humoral factors or altered cardiac gene expression lead to

cardiac hypertrophy, a fundamental adaptation of the adult heart [118]. The central role

of Ca2+ signaling in cardiac hypertrophy is apparent from studies on transgenic mice in

which the overexpression of genes encoding for components of this pathway, such as

2+ the L-type Ca channel, calcineurin and NFAT, TRD, JCT, CSQ2 and Gqα, all result in

hypertrophy [119].

Transgenic mice that overexpress CSQ2 have a particularly marked phenotype

that is characterized by severe cardiac hypertrophy and decreased depolarization-

induced Ca2+ release from the SR. The overexpression of CSQ resulted in a

compensatory decrease in the expression of its binding partners (JCT, TRD and RyR2)

within the RyR2 complex [106, 120]. The increase in transcription during hypertrophy

might result from subtle alterations in the spatio-temporal properties of the individual

Ca2+ spikes. Indeed, a broadening of the Ca2+ transient or an increase in its amplitude

have been recorded in cases in which hypertrophy is induced by modifying the levels of

27 proteins associated with RyR2 [119].

Table 1-2. Summary of junctional SR protein overexpression and suppression studies Gene Overexpression Animal Name Effect on Other Proteins Implications Ref. Symbol /Suppression Model Downregulation of TRDN Triadin Null Mouse --- [129] junctophilin-1 and -2 Hypertrophy, Reduced expression of JCT impaired relaxation Overexpression Mouse [130] and RyR and reduced contractility No changes in SERCA2a, CSQ, PLB and PMCA Mild cardiac Co- ATPase levels. RyR hypertrophy, overexpression expression levels prolonged Mouse [131] of canine TRD decreased 28%, however relaxation, a and canine JCT the phosphorylation of blunted response to S2809RyR was increased by β-adrenergic 57% Reduced expression of RyR2 (50%), CSQ2 Loss of Ca2+ (~60%), JCT (~92%) and release units, junctophilin-1 and -2 Null Mouse impaired E-C [132] (~40%). No significant coupling, and changes in the non- cardiac arrhythmias junctional SR proteins: SERCA2a, PLB Abnormalities in muscle contraction, narrowing of junctional SR, JCTN Junctin Overexpression Mouse --- [133] compaction of CSQ and excess T- tubule SR interactions Increased Overexpression Mouse --- susceptibility to [134] atrial fibrillation Enhanced cardiac No significant changes in function in vivo, RyR, phosphorylation of JCT deficient S2809RyR, TRD or CSQ, cardiomyocytes Null Mouse 2+ [135] histidine-rich Ca -binding exhibited increased protein, FKBP12.6, SERCA contractile and or phospholamban levels Ca2+ cycling parameters

Calseq Hypertrophy and Murine CSQ No downregulation of RyR, impaired Ca2+ - CASQ - Mouse 2+ [136] overexpression TRD or JCT induced Ca - uestrin release

Impairment in Ca2+ -induced Ca2+ - release and Canine CSQ2 Reduced expression of [106, Mouse decreased Ca2+ RyR, TRD and JCT overexpression spark frequency, 137] severe hypertrophy and heart failure

28

Ablation of the CASQ2 gene in mice (CASQ-/-) produces a cardiomyocyte

phenotype in which the SR Ca2+ release is not significantly different from that of wild-

type myocytes. A 50% increase in total SR volume in these mice could contribute to the

lack of change in the free Ca2+ concentration. Another possible explanation would be

that an increase in other SR Ca2+ binding proteins has occurred in response to the loss

of the Casq2 gene [121]. Subsequent studies of a heterozygote (CASQ+/-) mouse

models exhibited 25% reduction in CSQ2 levels with no effects on other junctional SR

proteins. The CASQ+/- model showed spontaneous SR Ca2+ releases, due to an

increase in diastolic SR Ca2+ leak that was similar to changes seen in some CPVT

models [122] that is; dysregulation of Ca2+ release.

Thus, mice with chronic changes in CSQ2 appear to develop general

malformation of Ca2+-release complexes. Functional changes reported in these mice,

however, may not be useful models for deciphering the functions of cardiac CSQ2.

Nevertheless, studies of heterozygote mice (CASQ2+/-) that exhibited roughly 25%

reductions in CSQ2 levels may be useful since the generation of a so-called “diastolic

leak” supports a function for CSQ2 in regulating the closing of the RyR2 channel. A

significant number of point mutations in human CASQ2 are also associated with a

recessive form of CPVT [123, 124], and studies have suggested similar clinical phenotypes for RyR2- and CSQ2-induced diseases [125]. Moreover, partial knockdown

of CSQ2 [122] also leads to a phenotype similar to CPVT, supporting a major role for

CSQ2 in regulation of RyR. In depth studies of the first reported mutation in CSQ2

(D307H) showed that the mutant protein was not capable of undergoing the appropriate

conformational changes for the Ca2+ concentrations expected to occur inside junctional

SR lumens, nor was it capable of binding efficiently to its target proteins JCT and

29

TRD[126].

1.7 Cytoskeleton

1.7.1 Cardiac cytoskeletal components

The integrity of cardiomyocytes is maintained by the cytoskeleton, which is divided morphologically into nuclear, membrane-submembrane, sarcomeric and extra- sarcomeric [127]. The nucleus is supported by a nuclear lamina, located below the inner nuclear membrane and consisting of a dense network of intermediate filaments, lamins, and interacts with both chromatin and the nuclear envelope. This whole complex is connected to the extra-sarcomeric cytoskeleton, actin, and T-tubules [128].

The membrane-submembrane cytoskeleton serves as the connection between sarcomeres and sarcolemma, and consists of the focal adhesion proteins (vinculin, and integrin), spectrins (spectrins, ankyrin and 4.1 proteins) and the dystrophin- dystroglycan complexes [138].

The sarcomeric cytoskeleton consists of thin filaments (cardiac beta- and gamma-actin, C- I- and T-troponins and alpha-tropomyosin), thick-filaments (myosin and myosin binding proteins C, H and X) and the Z-disk. The sarcomere, extra- sarcomeric cytoskeleton and the sarcolemma meet at the Z-disk. The extra-sarcomeric

cytoskeleton is composed MTs, intermediate desmin filaments and actin microfilaments

[138].

1.7.2 Microtubules and endoplasmic reticulum

Electron microscopy shows a close relationship between ER tubules and MTs,

where they are closely parallel over considerable distances [139, 140]. Furthermore,

evidence from studies utilizing live cell imaging show that MTs drive ER tubule

extension. Tubule extension is mechanistically achieved in three distinct ways:

30 association between ER and the (+) end of MTs through tip attachment complexes

(TAC), sliding of ER membrane on MTs that move through the cell as an intact entity,

and a more predominant and faster motor driven movement [141-143].

In vitro formation of ER tubules and network can be generated without the

cytoskeleton [144], thus, MTs are essential more in ER tubule extension and movement

than formation [145]. In vivo, stable interactions between MTs and ER associated

proteins are important for the ER network [146].

1.7.2.1 Microtubules

MTs are hollow cylindrical structures (diameter 24 nm), built from 13 parallel

protofilaments. They are composed of repeated heterodimers of 55 kDa α- and β- tubulin globular proteins. α- and β-tubulin molecules are homologous but not identical, each comprises several isoforms and, thus can have distinct locations in the cell, and perform different functions [127]. MTs are dynamic where they undergo phases of growing and shrinking. The binding and hydrolysis of GTP by tubulin subunits is known to lead to dynamic instability of the MTs [147].

MTs form a network that spans transversely and longitudinally along the cardiomyocyte axis and densely around and between nuclei [140, 148]. More generally,

MTs play a role in maintaining cell integrity [149] and are involved in organelle transport

[150], protein synthesis, and intracellular transport [151, 152]. Studies on membrane ion channels in cardiomyocytes support the role of MTs in ion channel delivery [153,

154].

1.7.2.1.1 Microtubule disrupting agents

MT depolymerizing agents cause a drastic alteration in the morphology of the

ER. Studies have demonstrated that many intracellular events rely on proper ER

31 network morphology. Mutations in proteins that localize to ER, identified in a class of

neurological disorders underscore the medical importance of understanding intracellular

membrane protein trafficking and localization that allow the ER to serve its

multifunctional roles.

Nocodazole (methyl [5-(2-thienylcarbonyl)-1H-benzimidazol-2-yl]) is a synthetic

compound that depolymerizes MTs and inhibits tubulin self-assembly [155].

Nocodazole acts through inhibition of polymerization of tubulin subunits, leading to de-

polymerization of dynamic microtubules [156, 157]. ER network collapse towards the

center of the cell and the formation of sheet-like structures has been demonstrated

upon nocodazole depolymerization of the MTs [158, 159].

1.8 Adenovirus mediated overexpression in cardiomyocytes

Several key characteristics of neonatal and fetal cardiac myocytes such as the

dynamic culturing nature, expression of signaling molecules, transcription factors, Ca2+ handling protein isoforms and morphological and sarcomeric organization have made difficult the extrapolation of data to adult cardiac myocytes [160-162]. Therefore, in vitro gene transfer through viral vectors to isolated adult cardiomyocytes is a powerful experimental approach in understanding structure and function.

Adenovirus is a member of the Adenoviridae class of viruses. The Ad vectors are one of the most efficient methods for in vivo and in vitro transduction. The most commonly used biomedical reagents are engineered replication-deficient serotypes Ad

2 and 5, where CMV promoter drives foreign gene expression. The adenovirus virion consists of a proteinacous capsid that surrounds the double-stranded DNA, and vertices that serve to attach the virus to the host cell membrane containing coxsackievirus

(CAR) or adenovirus receptors [163]. Once the Ad is endocytosed, the double stranded

32

DNA migrates to the nucleus and is transcribed [164].

In the studies described in Chapter 4 and 5, expression of canine junctional SR

proteins was performed at comparable levels to endogenous proteins in cultured adult

rat cardiomyocytes using Ad vectors, driven by CMV promoter.

33

CHAPTER 2

MATERIALS AND METHODS

2.1 Animals and heart tissue

The investigation conforms to the Guide for the Care and Use of Laboratory

Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996). The studies performed using Sprague-Dawley (Rattus norvegicus) rats

(Harlan Laboratories, IN) were approved by the Wayne State University Animal

Investigation Committee (Appendix B).

Heart tissue was obtained from sham-treated control dogs (C) and dogs in heart failure (HF) as frozen samples saved in liquid N2. Ischemia-induced HF (HFi) was

induced by multiple sequential intracoronary embolization’s with microspheres performed in mongrel dogs 1-3 weeks apart over 6 months, and was discontinued when left ventricular (LV) ejection fraction was ≤35%, as previously described [165], and animals were sacrificed after 3 additional weeks. Samples of the posterior wall from the

LV were stored at -80 oC, and thawed prior to tissue homogenization. For tachypacing-

induced HF (HFt), LV dysfunction was induced by rapid ventricular pacing in mongrel

dogs at >180 beats/min for >4 weeks to produce HF (ejection fraction ≤ 35%), and

animals were generally sacrificed within 1-3 weeks [166]. Samples of LV posterior wall

and septum were removed, and stored in liquid N2. Samples of frozen tissue used in

the present studies were thawed prior to tissue homogenization.

2.2 Purification of CSQ2

CSQ2 was purified from frozen tissue samples (2-5 g) as previously described

[167, 168]. Briefly, heart tissue samples were homogenized using Polytron

homogenizer (PT 10-35) at speed 5 for 30 sec in a Buffer A pH 8.0 containing 15 mM

34

Tris Base, 200 mM NaCl, 0.5 mM EGTA and a phosphatase inhibitors mixture of 5 mM

NaF, 5 mM Glycerol 2-phosphate Disodium salt Hydrate and 5 mM NaPPi, were added

to the homogenate. Triton X-100 was then added to the homogenate at 1%.

Homogenates were then transferred to centrifuge tubes and centrifuged using Sorvall

RC 5B PLUS (SS-34 rotor) at 35,000 x g for 45 min after which supernatants were

collected.

The collected supernatant was then applied to an anion exchange

chromatography column of DEAE-sephacel (Diethylaminoethyl Sephacel© (Sigma

I6505). The column was then washed with several volumes of Buffer A pH 8.0

containing 220 mM NaCl and 0.5% Triton X-100 then followed by Buffer A pH 8.0

containing 220 mM NaCl. A salt dependent elution was then performed with Buffer A

pH 8.0 containing 700 mM NaCl (Buffer B).

The collected elutions were then applied to a column of phenyl-Sepharose CL-4B

(Sigma P7892) equilibrated with Buffer B pH 8.0. The column was then washed with

several volumes of Buffer B to remove contaminants. A Ca2+ dependent elution was

then performed using Buffer B with 20 mM CaCl2. Spectrophotometry was performed at

280 nm on the collected elution fractions. The protein samples were then desalted by

repeated washes with HPLC grade water (Fisher Scientific) through centrifugal filters

(Amicon® Ultra-0.5 ml 10K, Millipore UFC501096). Concentrated fractions (1 g) was

run on acrylamide gel and visualized using Coomassie® Blue stain on gel (section

2.5.2).

2.3 Cardiomyocyte isolation

2.3.1 Surgery

Hearts were obtained from male Sprague Dawley rats weighing between 175-

35

200 g. Following standard protocol the animals were anesthetized with 300 mg/kg bodyweight by peritoneal injection of 5% Avertin (tribromoethanol) working solution in

PBS (Sigma, P4417). The working solution was prepared from a 50% stock of 2,2,2-

tribromoethanol (Sigma T48402) dissolved in 2-methyl-2-2butanol (Sigma 240486).

The incision area was sterilized using 70% ethanol. The chest was opened and the

heart was excised with a large portion of the aorta intact and placed immediately in cold

solution A (described In section 2.3.2).

2.3.2 Isolation protocol

Cardiomyocytes were isolated by retrograde perfusion using the Langendorff

apparatus [169]. Briefly, the heart was hung with the aorta clamped and tied with a

suture to a 1.8 mm aorta cannula (Radnoti 140153-16), to ensure good perfusion the

aorta was not pushed too far into the heart. Perfusion with warmed/oxygenated Ca2+-

free solution A containing 10.0 mM HEPES (Sigma H-4034), 1.2 mM KH2PO4, 5.0 mM

Pyruvic acid (Sigma P5280), 110.0 mM NaCl (Sigma S6191), 1.2 mM MgSO4 (Sigma

M2643), 5.0 mM creatine (Sigma C3630), 4.8 mM KCl (Sigma P5405), 11.0 mM

Glucose (Sigma G7528) and 5.0 mM taurine (Sigma T8691) was carried out for 5 min. to wash out the heart. Enzymatic dissociation was then carried out using solution A with

5 mg Liberase TM Research Grade (Roche 05401127001) and 0.1 mM CaCl2 (Sigma

C7902). After 15-20 min of enzymatic digestion, the ventricles were cut from the

cannula and placed in a dish along with the enzyme containing solution A in a 37 oC water bath for 10 min for further enzymatic digestion. The mixture was then filtered through nylon mesh into a 50 ml Falcon conical tube. The filtrate was spun in swinging bucket centrifuge for 2 - 5 min at ~1200 rpm until a light pellet formed. The cell pellet was resuspended with solution A containing 250 μM CaCl2, centrifuged for 2 - 5 min at

36

~1200 rpm until a light pellet formed, supernatant was discarded and cell pellet was

resuspended with 500 μM CaCl2 centrifuged for 2 - 5 min at ~1200 rpm until a light pellet formed, supernatant was discarded and cell pellet was resuspended as described below (section 2.3.3).

2.3.3 Cardiomyocyte culture

Cells were resuspended in Medium 199 (Invitrogen 12340-030) containing 2% bovine serum albumin (Sigma A1933), 2 mM carnitine (Sigma C0283), 5 mM creatine

(Sigma C3630), 5 mM taurine (Sigma T8691), 2 mM L-glutamine, 2 mM Glutamax-1

(Invitrogen 35050-061), 25 M blebbistatin (Toronto Research Chemicals B592500), 1%

Penicillin-Streptomycin (Invitrogen 15070-063), 1% ITS (Sigma I3146), 2 mM L- glutamine (Sigma G7513), 1% MEM non-essential amino acids (Invitrogen 11140-050).

The cells were examined under the microscope (Nikon Eclipse TS100) and then stained with trypan blue (0.4%) (Sigma T8154) and counted using hemacytometer (Reichert) for plating. The isolated cardiomyocytes were plated in 6-well dishes (50,000 cells/well) containing coverslips coated with laminin (Sigma L2020) at 37 °C with 5% CO2 and cultured up to 48 h.

2.4 Cell treatments

2.4.1 Adenovirus infection

Adenoviral CSQ2-DsRed (Ad.CSQ2-DsRed), CSQ2 (Ad.CSQ2), TRD (Ad.TRD) and JCT (Ad.JCT) vectors were previously described [84, 170, 171]. To determine the virus concentration (plaque forming units, pfu/l), HEK 293 cells (ATCC) were plated at

0.2 x 106 cells/well and the viruses were added the next day as follows: 10, 3, 1, 0.3,

0.1, 0.03, and 0 µl. For the purposes of estimating a concentration we assumed that

100% decimation of the cells was a multiplicity of infection (MOI) of 1. This assumption

37 was frequently checked by commercial assays. Calculation of pfu/cell was determined based on viable rod shaped cardiomyocytes not total cell count.

As previously described [19], adenoviruses were added directly to dishes 2 h

post-plating at MOI=200 (1x). In Chapter 5, the cardiomyocytes were infected with up to

5x Ad.CSQ2 in order to visualize the initial sites of accumulation following its protein

synthesis. For all adenoviral studies, treatments were carried out for 12 or 16, 24, 30

and 48 h as indicated before harvesting for biochemical analysis and fixing of coverslips

for microscopy.

2.4.2 Nocodazole treatment

Nocodazole (Sigma M1404) (20 M) was diluted in Medium 199 and added to

cardiomyocytes 16 h post-plating. Cardiomyocytes treated (+) with nocodazole up till 32 h in culture where as cells cultured under similar conditions without nocodazole (-),

served as control. Cells were fixed for immunofluorescence studies after 48 h in

culture. Treatments with nocodazole for last 2 h prior to harvesting and fixation,

however, did not show any marked disruption. Therefore, we opted to add nocodazole

12-16 h post-plating (Appendix C Figure A-3).

2.5 Electrophoresis

2.5.1 Protein extraction and concentration determination

Protein extraction from cultured cardiomyocytes was performed as follows,

immediately after removal of coverslips for immunofluorescence analysis (see section

2.6.3.1), 6-well dishes were placed on ice and cells were scrapped and transferred with

media into 15 ml Falcon conical tubes. The cells were centrifuged using ADAMS

DYNAC centrifuge (Clay Adams Inc.) for 10 min at 4 oC and media was discarded. The

cell pellet was resuspended with PBS and centrifuged at the same settings.

38

Supernatant was again discarded and cell pellets were resuspended with 200 µl extraction buffer (10 mM Tris, 200 mM NaCl, 1% triton X-100, 0.5 mM EGTA, pH 8.0),

centrifuged after which protein extracts were collected and saved in -80 oC for later

determination of protein concentration by Lowry assay [172] and electrophoretic

analysis.

2.5.2 Gel electrophoresis

SDS-PAGE was carried out according to Laemmli [173] on 7% or 10%

acrylamide gels using Hoefer® GE600 Chrome apparatus (Amersham) or on 10% Bis-

Tris MIDI gels (Invitrogen) according to manufacturer. Gels were transferred to

nitrocellulose membranes (0.45 m, BioRad Laboratories) and stained with Amido Black

(Sigma).

2.5.3 Concanavalin A binding to glycoproteins

Similar amounts of purified CSQ2 from heart samples were analyzed by

immunoblotting based on immunoblot quantification. Duplicate nitrocellulose strips

were then analyzed by immunoblotting (section 2.6.2) or Con A binding. For Con A

binding to CSQ2, one strip without prior non-specific protein blocking of membrane was

incubated in phosphate buffer saline with 0.2% tween20 (PBS/tween) containing Con A

conjugated to peroxidase (Sigma, L6397) at 10 μg/ ml for 90 min at room temperature

[174]. The strip was rinsed three times for 5 min each, and then developed using

enhanced chemiluminesence (ECL) reagents (Pierce 32106). ECL and quantitation of

digitized scans were performed to determine Con A binding/μg CSQ2 using ImageJ

(Wayne Rasband, National Institutes of Health).

39

2.6 Immunological methods

2.6.1 Antibodies

Antibodies used in immunoblotting and immunofluorescence assays were as follows: anti-CSQrat (Abcam ab3516), anti-CSQdog, anti-JCTdog and anti-JCTrat antibodies

were the generous gift of Dr. Larry Jones, Indiana University School of Medicine.

Monoclonal anti-CSQ2 antibodies used for assays in Chapter 2 are previously

described [85]. Anti-TRDdog antibodies were as previously described [175], anti-TRDrat were purchased from Thermo (MA3-927). Tubulin antibodies were as follows: anti- alpha tubulin (Sigma T6074) and anti-beta tubulin (Sigma T5201). Alexa Fluor 488 or

568 conjugated goat anti-rabbit IgG, Alexa Fluor 488 or 568-conjugated goat anti- mouse IgG secondary antibodies were purchased from Invitrogen. Relative specificity/selectivity of the antibodies used towards dog or rat junctional SR proteins indicated by (+) or (-) are shown in Table 2-1.

Table 2-1. Relative reactivity of junctional SR protein antibodies to dog or rat forms.

Species Reactivity Protein Target Antibody Dog Rat

anti-TRDdog + - polyclonal TRD anti-TRDrat - +monoclonal

anti-JCTdog ++ - monoclonal JCT anti-JCTrat + ++polyclonal

anti-CSQdog ++ - monoclonal CSQ anti-CSQrat + ++polyclonal

2.6.2 Immunoblot

Immunoblotting was carried out as previously described [176]. Briefly, the nitrocellulose membranes were blocked with 1 mg/ml bovine serum albumin in

40

PBS/tween for 15 min and then incubated with primary antibodies solution in PBS/tween for 90 min and then washed three times with 15 min. Primary antibody dilutions were adjusted to assure that ECL signals for multiple antigens on a single exposure of film remained within a linear range. Membranes were then incubated in PBS/tween with horseradish peroxidase-conjugated goat anti-rabbit IgG and goat anti-mouse IgG secondary antibodies (1:30,000) (Jackson ImmunoResearch Laboratories) for 45 min and then washed with PBS/tween. ECL reagent was used to detect immune complexes, which were then visualized by autoradiography using high resolution

Amersham Hyperfilm ECL film (GE Healthcare). Autoradiograms were quantified using

ImageJ.

2.6.3 Immunofluorescence

2.6.3.1 Immunocytochemistry on isolated cardiomyocytes

Briefly, cells grown on coverslips were fixed with 4% paraformaldehyde in PBS for 15 min and then permeabilized for 10 min in 0.2% PBS/tween. Coverslips were blocked in PBS with 0.2% Tween-20 (PBS/Tween) and 2% goat serum at room temperature for 20 min. Cells were then incubated in PBS/Tween with primary antibody

(1:100) overnight at 4°C, followed by washing in PBS/Tween, and incubation with goat anti-rabbit IgG or goat anti-mouse IgG antibodies conjugated to Alexa Fluor dyes (1:100 dilution in PBS/Tween for 120 min). Cells were counterstained with NucBlue Fixed Cell

ReadyProbes Reagent (Molecular probes R37606) for 30 min. rinsed, and mounted to microscope slides with ProLong Antifade Mounting Kit (Molecular Probes).

2.6.3.2 Immunohistochemistry on tissue sections

Frozen LV tissue from C and HF hearts were removed from liquid N2 and

embedded in OCT compound, sliced by cryostat (-21 °C, 10 m thickness), and dried

41 on Superfrost® Plus Gold slides (Fisherbrand). The slices were hydrated in PBS and the OCT washed three times in PBS at RT. Fixation was performed with 4% paraformaldehyde in PBS for 15 min, and washed three times, then blocked with 1% goat serum in PBS-tween for 4 h; all carried out at ambient temperatures. Antibodies were premixed before use to attain uniform distributions. The primary antibody mixture contained 1:50 dilutions of mouse anti-CSQ2 monoclonal (mAb) 4E7 and rabbit anti-

CSQ2 polyclonal (pAb), and was applied for 24 h at 4 °C. Equal volumes of the two antibodies contained roughly equivalent ECL signals against purified canine CSQ2 using the same respective anti-IgG goat antibodies used for immunoblotting. Slides were than washed three times for 5 min. The secondary antibody mixture contained

1:200 dilutions of goat anti-mouse Alexa Fluor 488 (Invitrogen) and goat anti-rabbit

Alexa Fluor 568, and was applied 4h at RT. Slides were then washed three times for 15 min in PBS/tween, RT, stained with 100 µM 4′-6-diamidino-2-phenylindole (DAPI) for 2 min, and washed for 10 min in PBS/tween, RT. Coverslips were mounted for imaging using a drop of ProLong Gold Antifade (Invitrogen).

2.7 Fluorescence microscopy

Imaging of tissue and cardiomyocytes was performed using a 63 ×/1.4 oil objective on a Leica DMI 6000B system. All confocal images were acquired at the same settings and were concurrently processed and optimized offline for publication using Photoshop (Adobe Systems Inc.). Fluorescence intensity quantification was performed using ImageJ (Wayne Rasband, National Institutes of Health).

Transverse lines of 50 pixels were used to select areas that extended from the cell edges and included the center of one nucleus (in tissue staining) and to the edge of the nucleus (in cell immunostaining). The intensity of each color channel was plotted

42

(red: pAB-CSQ-2, green: mAB-CSQ-2, blue: DAPI) in consecutive 3-μm steps distance from the nucleus. Values are plotted as fractional deviations from the average fluorescence intensities for each channel. Channel values were standardized for plotting using Microsoft Excel.

2.8 Quantitative PCR

RNA extraction from heart samples was performed using RNeasy Fibrous tissue

Mini Kit (Qiagen). One microgram of DNase-treated total RNA sample was reverse

transcribed by reverse transcription (Applied Biosystems). Quantitative PCR with fast

SYBR green (Applied Biosystems Step One with v2.0 software) was performed using

canine specific primers designed to span one intron. Canine primers were as follows:

GAPDH: forward, CCCCTTCATTGATCTCAACTACATGGT reverse, CCCGTTGATGACAAGTTTCCCGTTC-TC CSQ2: forward, CCAAGCTTGCCAAGAA-GCTGGGT reverse, ACGTCAGCTGCAAACT-CGCCA ANF: forward, AGAGCGGACTGGGC-TGCAACA reverse, CCCAAGGGACGAGGGT-CTCTCA BNP: forward, ACTACGCAGCCCC-AAGATG reverse, CCTCAGCACTTTGCAGC-CCA

Relative quantitation method (∆Ct) was used to evaluate mRNA levels for CSQ2,

along with hypertrophic markers atrial natriuretic factor (ANF) and brain natriuretic

peptide (BNP). Fold change (2-ΔCt) relative to glyceraldehyde 3-phosphate

dehydrogenase (GAPDH) mRNA levels was used. GAPDH mRNA levels were highly

similar between C and HF samples.

2.9 Mass spectrometry

In order to measure the mass to charge ratio (m/z) mass spectrometry was used.

Small highly charged droplets are formed by electrostatic dispersion of protein solution

through a glass capillary subjected to a high electric field. In order to calculate the molecular weight of a particular peptide, the observed m/z ratio has to be corrected for

43

(multiplied by) the number of attached protons [177]. Electro spray ionization mass

spectrometry of purified CSQ2 (ESMS) was performed at the Keck Biotechnology

Laboratories, Yale University, New Haven, CN using a Q-ToF mass spectrometer

(Micromass, Altrincham, UK), as previously described [167]. Protein samples were

resuspended or diluted in 20% acetonitrile, 0.1% formic acid. The proteins were

trapped on a Michrom (Auburn, CA) protein cap trap using a Waters (Milford, MA)

CapLC. Trapped proteins were eluted into a Waters Q-Tof Ultima mass spectrometer

and their masses deconvoluted using the MaxEnt1 algorithm.

2.10 Statistical analysis

Data are presented as mean ± SEM. To determine differences between C and

HF groups, Student’s t-tests were performed, and P<0.05 was considered statistically

significant. Microscopy data for tissue slices from two hearts as mean ± SD.

44

CHAPTER 3

ALTERED CALSEQUESTRIN GLYCAN PROCESSING IS COMMON TO DIVERSE

MODELS OF CANINE HEART FAILURE

3.1 Rationale

The N-linked glycan in CSQ2 from mammalian heart tissue normally undergoes a

high degree of trimming by intracellular mannosidases. Minor differences in CSQ2

glycan structures have been previously noted in larger mongrel dog hearts [167]

compared to that of purebred beagles [178]. Interestingly, in beagles, CSQ2 glycans

exhibit a dramatic reduction in mannose trimming following a 6-8 week period of

tachypacing that produces HF [178]. To determine whether this is an effect specific to

the smaller animal or to a single type of cardiac stress, we broadened such investigations to include two diverse experimental models of HF in the mongrel dog.

We report here that whether dogs receive ventricular tachypacing or are placed into failure by ischemia due to recurring coronary microembolizations, dramatic and qualitatively similar alterations of CSQ2 processing occur wherein normal CSQ2 is replaced in HF tissue with protein that has not undergone the same degree of posttranslational processing.

Hypothesis: CSQ2 trafficking within the cardiomyocytes is dramatically altered in mongrel canine populations using two of the leading models of heart failure induction

3.2 Results

3.2.1 Analysis of CSQ2 glycoforms and phosphoforms

Purified CSQ2 analyzed using ESMS resolves CSQ2 into its polymorphic spectrum of masses and provides quantification of individual CSQ2 molecular

structures. Detailed knowledge regarding mannose trimming of its 316Asn-linked glycan

45 can be deciphered from these mass spectra, which undergo loss of mass in 162 Da steps that are indicative of mannose trimming [167]. Superimposed upon the mass change for each glycoform can be additional mass changes due to changes in CSQ2 phosphorylation on its C-terminus (378,382,386Ser). The exact state of phosphorylation for a given glycoform is obscured by the coincidental overlap in mass changes resulting from mannose trimming (Δmass = 162 Da) and that resulting from differences from two

phosphates (2 x 81 Da = 162 Da) [167, 179]. Nonetheless, relatively large and very consistent differences in CSQ2 mass peaks were found between control and failed heart tissue samples occurring in steps of 162 Da and 81 Da that reflect differences in steady-state levels of these two ongoing posttranslational processes.

Purified CSQ2 from C mongrel dogs exhibited a range of molecular masses,

having a weighted average of 46,306 Da, a value that is 1037  41 greater than its

deduced molecular weight of 42,269 (Table 3-1). CSQ2 masses, 90% of which were

between 46,001 and 46,487 Da, corresponded to CSQ2 molecules having glycans that

have undergone processing by intracellular mannosidase(s) to structures

GlcNAc2Man3-5 (Figures 3-1 & 3-2). In contrast to the single uniform peak of CSQ2

masses present in C hearts, every heart from HF dogs contained CSQ2 molecules that

were less processed, and so were of greater mass. More striking was the appearance

of two broad groupings of mass peaks, demarcated Pool I and II (Figure 3-1). A

prominent lower mass pool in HFt and HFi (Pool I) was relatively close in mass to the

single pool of CSQ2 found in C hearts, yet larger by 274  94 and 326  85 Da,

respectively (Table 3-I). The highly processed CSQ2 molecules found in all canine

hearts reflects the successive removal of mannose in more distal subcellular

compartments, processed by mannosidase(s) beyond that of the early ER [167].

46

Although not structurally identical in C, HFi, and HFt hearts, it is hypothesized that this larger pool of highly processed protein in each heart corresponds to junctional SR

CSQ2.

Table 3-1. Molecular masses (weighted averages ± S.D.) of CSQ2 from canine heart tissue samples (Da)

Control HF (Tachypacing) HF (Ischemic)

Relative protein amounts Pool I (Man1-Man7 glycoforms) 100 68  20 68  9 Pool II (Man8,Man9 glycoforms) 0 32 32

Weighted mass averages All forms 46,306  41 46,811  116 46,798  196 Pool I (Man1-Man7 glycoforms) 46,306  41 46,633  85 46,581  94

Pool I (Increase from Control) N/A ∆AVG = 274  94 ∆AVG = 326  85

Pool II (Man8,Man9 glycoforms) N/A 47,290  88 47,193  30

The second broad grouping of mass peaks (Pool II, Figure 3-1 and Table 3-I)

was comprised of protein that underwent either no mannose trimming, or removal of a

single mannose. This pool of CSQ2 is likely only acted upon by the well-characterized

ER α-1,2 mannosidase, characteristic of early ER, and known to produce Man8 from

Man9 [180, 181]. This is the mannose trimming typical of CSQ2 expressed in every non-muscle cell examined to date [182]. This higher-mass pool of CSQ2 molecules was only present in HF tissue, entirely absent form C mongrel dog heart tissue.

47

Figure 3-1. CSQ2 masses are altered in heart tissue from mongrel dogs in heart failure. a) CSQ2 was purified from control (C) canine heart, and hearts induced into failure by repeated microembolisms over 6-9 months (HFi) or by tachypacing (HFt) for 4-8 weeks. CSQ2 was purified from frozen tissue samples and intact CSQ2 was analyzed by ESMS. Differences in CSQ2 mass isoform peaks correspond to the successive loss of mannose sugars (mannose trimming, ∆mass = 162 Da) from the CSQ2 glycan. Superimposed on each glycoform are changes in mass brought about by varying numbers (0-3) of phosphates on CK2-sensitive sits (∆mass = 81 Da). Masses and mannose content are shown across the bottom and top, respectively. CSQ2 mass peaks corresponding to Man9,8 glycoforms are shaded. b) Summary raw data for all 15 heart samples (12 dog hearts). Mass peaks corresponding to glycoforms and phosphoforms are plotted as a percentage of the total (%CSQ2 molecules). Five C hearts (green spectra), 3 HFi hearts (red spectra), 4 HFt hearts (blue, posterior wall, lavender, septal wall). Shaded area designates rough ER with Man9,8 (Pool II), while further mannose trimming is designated as junctional SR (and Pool I). The presumed mass of CSQ2 following biosynthesis and folding (mass = 47,478 Da; Man9 with 3 phosphates) indicated by an asterisk (*).

Redistribution of CSQ2 molecules from C heart tissue into two mass pools of

Man1-7 (Pool I) and Man9,8 (Pool II), was further analyzed by calculating separate

Gaussian distributions based upon the averages calculated from the three cohorts of

48 mongrel dogs (Figure 3-2 panel a). Here we compared the average molecular masses

for the two putative CSQ2 pools, along with their relative abundances (relative peak

heights), and their standard deviations (relative peak width). Results from the current

three cohorts of mongrel dog heart samples were also compared with C and HF cohorts

previously reported for beagles [178]. From these canine heart samples, a general

pattern emerges wherein CSQ2 from C heart tissue (Figure 3-2 panel a, green curves)

is consistently comprised of a more highly processed collection of molecules, whereas

HF tissue has degraded those molecules, and replaced them with less processed, or

post-translationally "younger", molecules that accumulate in two distinct states of

posttranslational processing (Figure 3-2 panel a, red curves).

Because mannose trimming occurs in successive steps, and reflects protein

biosynthesis, processing, and degradation, the percentage of total CSQ2 remaining at

each step provides a perspective of its overall turnover (Figure 3-2 panel c). In C dogs,

trimming of the first 4 mannoses was not accompanied by any retention, and so 100%

of CSQ2 molecules proceed to latter steps of mannose trimming. The first four steps of

processing correspond to cleavage of (1-2) linkages by an ER (1-2)-mannosidase

(Figure 3-2 panel b, green circles). CSQ2 accumulated in the form of molecules with

additional mannose trimming, with successively fewer molecules of CSQ2 trafficking

forward. This analysis suggests that the actions of a second enzyme ((1-3)-

mannosidase) might therefore be associated with the subsequent loss of mannose,

leading to loss of CSQ2 from the cardiomyocyte. We cannot determine from the mass

spectrometric data whether all molecules eventually underwent trimming to Man 3 on

their path to degradation, or whether all molecules with glycans of Man5-3 were

degraded directly.

49

Figure 3-2. Further evaluations of CSQ2 mass spectra data. For the analyses shown here, the direction of CSQ2 posttranslational processing is reversed from Figure 3-1, which reflected the orientation of mass spectrometric data. In panels a, b and d, CSQ2 processing occurs left to right a) Weighted averages of CSQ2 masses were calculated individually for Man9,8 and Man1-7 to construct a model of CSQ2 glycoforms based upon low or high levels of mannose trimming (cf. Table 3-1). Control mammalian hearts (C) show one peak of highly mannose-trimmed molecules (green peaks). In failing hearts (HFi and HFt), the green peak is no longer present, having apparently degraded. In its place is found a distinct phenotype exhibiting two CSQ2 pools (red peaks), consisting of a prominent pool (Pool II) of poorly processed (Man9,8) protein and a pool of junctional SR CSQ2 (Pool I) with glycan processing closer to that of control hearts. Similarly modeled are data previously reported in tachypacing of beagle hearts that led to HF (Ctb/HFtb) [178]. CSQ2 expressed in non muscle cells [182] produces only ER- type (Man9,8) CSQ2 (bottom panel) b) Sample structures of CSQ2 protein molecules (C-termini) corresponding to masses as indicated. After biogenesis in rough ER, the N- linked glycan on 316Asn is -GlcNAc2Man9, whereas CK2-sensitive sites (378,382,386Ser) are fully phosphorylated, since phosphorylation occurs in the cytoplasm during translocation of the nascent polypeptide [183]. Moieties are GlcNAc (black boxes), mannoses (blue and green circles) -1,2 linked mannoses (green circles), phosphates (red circles). Mannoses and phosphates removed by mannosidases and phosphatases are represented by lightened symbols.

3.2.2 Analysis of CSQ2 glycans by Con A binding

To provide a gel-based assay to corroborate the mass spectrometric analysis of

CSQ2 glycans, we measured Con A binding to purified CSQ2 on nitrocellulose transfers. Con A is a lectin known to bind to high mannose forms of N-linked glycans

[174, 184]. CSQ2 from hearts of HFt dogs, but not HFi dogs, bound significantly higher

50 levels of Con A than CSQ2 from C dogs, indicative of the accumulation of the

unprocessed protein (Figure 3-3 panels a, b). For tissue samples analyzed both by

ESMS and Con A binding, a non-linear correlation was found between the percent of

CSQ2 in Pool II (ER CSQ2) and Con A binding (Figure 3-3 panel c). This likely

reflected increased levels of Man9 compared to Man8, but this distinction was difficult to

confirm by ESMS because of the overlap of mass peaks due to overlap of masses

resulting from loss of mannose (mass = 162 Da) and phosphate (mass = 81 Da) [185].

Figure 3-3. Con A binding to purified CSQ2 from C and HF tissue samples. a) Con A binding to purified CSQ2 on nitrocellulose membranes. Measurements were carried out in 3 independent determinations of the same sample, from 16 hearts, including samples of HFt posterior wall and septum (LV regions). Averages for each animal cohort of C, HFi, and HFt are mean  SEM.; significance (p < 0.05) is indicated by asterisk (*). Values for LV regions are mean  SD. b) Sample autoradiograms of CSQ2 antibody binding (upper band) and Con A binding (lower band) for C and HFt sample. Data shown in panel a is Con A divided by CSQ2 antibody binding, normalized to the values obtained for CSQ2 from HEK cells (cf. Figures. 3-1a, 3-2a). c) Comparison of percentage of CSQ2 molecules in ER (Pool II glycans) plotted against Con A binding, for samples in which both types of data were obtained.

3.2.3 Steady-state CSQ2 protein and mRNA levels

Total steady-state levels of CSQ2 protein were measured in tissue homogenates from C, HFi, and HFt mongrel dogs. Decreased total levels of CSQ2 were found in both

51

HF dogs compared to C (Figure 3-4 panels a, b) with mean levels of CSQ2 reduced in

HFi and HFt by 86% and 58%, respectively, although neither was statistically significant.

Data were also obtained for CSQ2 mRNA levels in HFi and HFt. Compared to C hearts; mRNA in HFi and HFt was increased by 33% and 72%, respectively (Figure 3-4 panel c). Only changes in CSQ2 mRNA levels in HFt dogs attained statistical significance

(p=0.04).

Figure 3-4. Steady-state levels of CSQ2 protein, mRNA, and protein to mRNA ratios in canine hearts. Total numbers of dogs were: C, n=6; HFi, n=5; HFt, n=5. Samples from each dog were measured a minimum of four times. a) Canine heart homogenates were analyzed by immunoblotting (30 μg/lane). Representative immunoblot data: Amido black-stained nitrocellulose (Protein Stain) and autoradiogram for α -actinin and CSQ for C, HFi and HFt. Molecular weight markers (left margin) are in kDa. b) Summary data for C, HFi and HFt dogs normalized to α-actinin as loading control. Values are from 3 measurements for 5 dogs (mean  SEM, p-values for t-test, significance < 0.05 indicated by asterisk (*)). c) Data for levels of mRNA for CSQ2, ANF, and BNP, determined by quantitative PCR for C, HFi, and HFt dogs (mean  SEM, p values for t- test; significance < 0.05 indicated by asterisk (*)). Values on right correspond to ANF and BNP mRNA levels. d) CSQ2 protein to mRNA ratios calculated for each dog, and averaged for C, HFi, and HFt dogs (mean  SEM, p values for t-test; significance < 0.05 indicated by asterisk (*))

Transcript levels for ANF and BNP were also determined as a measure of hypertrophy in C, HFi, and HFt groups. Mean values for these two markers were highly

52 increased in both HFi (ANF, 16-fold; BNP, 51-fold) and HFt (ANF, 18-fold; BNP 96-fold)

tissues compared to C (Figure 3-4 panel c). The increase in dog HF samples, while very large, did not attain statistical significance, as large variations in transcript levels were measured for individual dogs. Reasons for variability were not further investigated.

Since protein levels showed a tendency to decrease, in spite of increased mRNA levels, we determined protein levels per mRNA levels, averaged for each dog (Figure 3-

4 panel d). For HFt dogs, this measure of protein/mRNA was reduced by 3.0-fold

(p=0.003), whereas a lesser but not significant reduction was found for HFi dogs

(p=0.08).

3.2.4 Immunohistochemical analysis of canine tissue

Given the roughly one-third of CSQ2 that accumulates in an unprocessed form in

cardiac HF tissue, we tested whether this separate mass pool of CSQ2 might be

apparent by immunohistofluorescence microscopy. The rough ER responsible for

CSQ2 biosynthesis is thought to localize to membranes surrounding myonuclei, based

upon studies of CSQ2-DsRed overexpression in cultured adult rat cardiomyocytes[117].

To determine whether there was enhanced CSQ2 immunofluorescence relative

to myonuclei in HF compared to C tissue, we analyzed thin sections of heart from C

tissue and from two different HFt tissue samples. HFt samples were chosen that

exhibited high levels of ER-CSQ2 by ESMS. Quantitative analysis of confocal

immunofluorescence from C and HFt tissue, showed no significant differences in

relative CSQ2 levels across the cardiomyocyte (Figure 3-5).

53

Figure 3-5. Detection of CSQ2 in canine tissue slices by immunofluorescence, and CSQ2 antibody reactivity. Tissue sections were analyzed by immunofluorescence using a mixture of polyclonal (poly, Red) and monoclonal (mono, Green) anti- CSQ2 antibodies, each diluted 1:50. Images were obtained as a series of optical slices (Z-stack), and single confocal images were selected for quantitation, as described in Methods a) Bar graphs show integrated intensities of fluorescence signals across a transverse width for 12 individual cells from five separate Z-stacks obtained for two different left ventricle tissue slices. Values are total fluorescent intensity over background (lowest intensity per channel), averaged for all 12 cells (mean intensity in each 3-μm window for red or green channel SD). Scale bar, 3 μm. b) Relative immunoreactivity for roughly equivalent IgG concentrations of the two antibodies. Lane 1, 1.0 μg purified canine CSQ2 from HEK cells, which contains ~1.5 mol phosphate on C-terminal sites and a glycan of structure -GlcNAcMan9,8 [167, 182]; lane 2, 1.0 μg purified of CSQ2 phosphorylation-site negative mutant Ser(378,382,386)Ala from HEK cells [183]; lane 3, 1.0 μg purified CSQ2 from Control canine ventricular tissue, which contains ~1.5 mol phosphate on C-terminal sites and an N-linked glycan of structure –GlcNAcMan4-6 ([179] and Figure 4). c) Sample confocal fluorescence images from C and HFt tissue slices. Scale bar 3 μm. Enlarged field (white inset) showing minor differences in immunoreactive localization. Scale bar 1 μm.

Interestingly, our data also supported the idea that there is more than a single

homogeneous pool of CSQ2 within canine cardiomyocytes. Commercial anti-CSQ

polyclonal antibodies produced the expected intense punctate immunofluorescence,

54 whereas canine-CSQ2-selective monoclonal antibody produced a similar but clearly distinct immunofluorescence (Figure 3-5 panel c). These differences in apparent subcellular localization were most likely due to differences in accessibility of the epitope

for the monoclonal antibody (264EFLEILKQV), not to selective posttranslational changes.

Immunoblot analyses of relative CSQ2 immunoreactivities for the two antibodies showed that neither the polyclonal nor monoclonal antibodies exhibited differences in

CSQ2 immunoreactivity that stemmed from its level of endogenous phosphate or its degree of glycan processing (Figure 3-5 panel b). This was tested by comparing reactivities against recombinant canine CSQ2 from HEK cells, in which little mannose trimming occurs [186], or a phosphorylation-site mutant Ser(378,382,386)Ala [183], or

CSQ2 from C dog heart which contains phosphate and undergoes substantial mannose trimming.

3.3 Summary

In this chapter, we have shown that the prevalent form of CSQ2 glycans in normal dogs is absent in HF dogs and is replaced with two new pools that accumulate

with different forms of the glycan. The most unprocessed form of CSQ2 accumulated to

as high as 40% of total CSQ2 in only weeks of the induced cardiomyopathy. The most

highly trimmed forms of CSQ2 were virtually absent in HF animals, indicating that CSQ2

was not processed as HF ensued and by the time the tissue was harvested (4 weeks or

greater) (Figure 3-2 panel a). Whether complete loss of this highly processed CSQ2 in failing hearts was due to normal attrition could not be determined from the data.

Nevertheless, in place of normal low-mass CSQ2, two higher-mass forms of CSQ2 accumulated in HF (Figure 3-2 panel a). Given the altered patterns of CSQ2 processing, and the reduction in CSQ2 protein levels, and the increase in mRNA levels

55

(Figure 3-3 panel b), the data infer an increased breakdown of CSQ2 in HF.

A better understanding of basic ER/SR protein traffic in adult cardiomyocytes is needed for interpreting the changes in CSQ2 we found in vivo. In Chapters 4 and 5, we therefore examined where within the cell are found newly formed junctional SR proteins.

56

CHAPTER 4

SORTING AND TRAFFICKING OF JUNCTIONAL SR TRANSMEMBRANE

PROTEINS ACROSS ER ALONG MICROTUBULES

4.1 Rationale

SR subdomains form at sites where protein complexes concentrate [26, 187]. In myocytes, atypical intracellular structure has prevented a clear understanding of the routes by which ER/SR compartments form and are dynamically maintained.

Relationships between the inner membrane subcompartments in cardiomyocytes have historically been more defined by ideas of Ca2+ cycling within the cell than by their cell

biological compartmentation or biogenesis. Studies do suggest that even at steady

state, the SR is a dynamic structure, and its motility is regulated by MTs and the

molecular motors dynein and kinesin (kif5b) [188]. Previous studies using the fusion

protein CSQ2-DsRed suggested that its biosynthesis was in or near the nuclear

envelope [116, 117].

To investigate cellular mechanisms for sorting and trafficking proteins to

junctional SR, we expressed canine forms of JCT or TRD in adult rat cardiomyocytes.

Protein accumulation over time was visualized by confocal fluorescence microscopy

using species-specific antibodies.

Hypothesis: Junctional SR transmembrane proteins TRD and JCT synthesis occurs in

juxtanuclear rough ER and traffics along ER and MT pathways to accumulate in

junctional SR

4.2 Results

4.2.1 Species-specific antibodies to junctional SR transmembrane proteins

To validate the relative immunoreactivities of antibodies used in this study, we

57 tested relative specificities by immunoblotting against samples from canine and rat heart microsomes. Anti-JCTdog and anti-TRDdog antibodies exhibited complete specificity to the canine forms (Table 2-1 and Figure 4-1). These antibodies would therefore permit the specific detection of acutely expressed canine junctional SR proteins from the steady-state levels of the native rat heart protein forms. Meanwhile, rat forms JCTrat and TRDrat can be detected using a second set of antibodies, although anti-JCTrat

detected both forms. We found minor species-specific differences in TRD mobilities but

not JCT (Figure 4-1 panel C).

Figure 4-1. Specificity of anti-TRD and anti-JCT antibodies. Samples (50 g) of canine heart microsomes (MVc), rat heart microsomes (MVr) and primary rat cardiomyocytes (HCr) were analyzed by SDS-PAGE (10% acrylamide) and transferred to nitrocellulose. (A) Amido black stained nitrocellulose, molecular weight markers in kDa. (B) Immunoblotting with antibodies anti-TRDrat and anti-TRDdog. (C) Immunoblotting with anti-JCTdog and anti-JCTrat. Molecular weight markers kDa, indicated on the left for each immunoblot.

4.2.2 Early accumulation of expressed JCTdog and TRDdog at juxtanuclear sites

To visualize the intracellular sites of earliest JCT and TRD accumulation and early trafficking, we analyzed cultured adult rat cardiomyocytes after 12, 24, 30 and 48h of overexpression. Confocal images of indirect immunofluorescence were generally

58 acquired as a Z-stack covering the height of the cell in contiguous series of optical slices of 0.5 m thickness, allowing us to most clearly determine ER/SR structures radiating outward from nuclei.

Figure 4-2. JCT accumulation at juxtanuclear sites Coverslips containing cardiomyocytes were fixed following no virus (-) or treatment with JCT (JCTdog) adenovirus for 16, 24, or 30 h, and remaining cells harvested for immune-blotting. Cells were double-labeled with anti-JCTdog (green) and anti-JCTrat (red) at each time point. Fluorescence images for cells were acquired using identical settings and processed offline at the same time, to permit direct comparisons. (A) Representative cells are shown for time points as indicated 16, 24, and 30 h, either for JCTdog the green channel (anti- JCTdog) alone, or merged with red JCTrat (anti-JCTrat). Panel c is the same image as shown in b following an arbitrary increase in the signal offline. Panels f and g are the merge images of d and e, RGB channels. Scale bar, 10 µm. (B) Magnification of outlined box (panel A-f) showing JCTdog appearing first at juxtanuclear sites. Scale bar 2 µm. (C) Immunoblot analysis of cardiomyocyte protein extracts showing JCTrat and JCTdog levels with increasing times of JCTdog overexpression. Immunoblot was performed using anti-JCTrat (detecting both native and expressed forms) and anti- JCTdog. Molecular weight marker on the left is 31 kDa.

JCTdog first appeared after 16-24 h of virus treatment, and its early distribution showed a restricted pattern of puncta that surrounded nuclei, when viewed in an optical plane through the center of a myonuclei. The most intensely fluorescent puncta were those that most closely encircled the nucleus (Figure 4-2 panel A). Additional puncta of lower intensity appeared to radiate away from the nuclear surface along transverse (Z-)

59

lines towards the periphery of the cell. JCTdog immunofluorescence co-localized with that of native JCTrat, validating its passage across, and concentration in, junctional SR.

With increasing incubation times, JCTdog accumulated at greater distances from the

nuclear surface, suggesting a peripherally directed movement of protein across the cell.

By 36-48 h of incubation, the distribution of JCTdog was qualitatively similar to that of

native JCTrat. The time course of JCTdog accumulation observed in the microscope was

validated by immunoblotting of harvested cells (Figure 4-2 panel C) JCTdog levels rose

significantly between 24 and 48 h, commensurate with the accumulation of fluorescent

puncta across the cell periphery.

TRDdog showed a pattern of accumulation that was similar to that of JCTdog.

TRDdog also first appeared in cultured cardiomyocytes around 16-24 h in culture (Figure

4-3). Newly synthesized TRDdog accumulated in cisternae that encircled nuclei at 16 h

when visualized in optical plane of the nucleus (Figure 4-3 panel A). TRDdog and TRDrat immunofluorescence co-localized and showed a very similar distribution (Figure 4-3 panels B, C) validating an apparent direct passage of TRDdog across, and concentration in, junctional SR. Thus, a common pathway and time course JCT and TRD accumulation across increasingly peripheral junctional SR puncta suggested sites of biosynthesis close to the cell surface, with an apparent direct membrane trafficking pathway transversely (radially) across junctional SR.

60

Figure 4-3. TRD accumulation at juxtanuclear sites. Coverslips containing cardiomyocytes were fixed following treatment with TRD (TRDdog) adenovirus for 16, 24, 30, or 48 h, and remaining cells harvested for immunoblotting. Cells were double- labeled with anti-TRDdog (green) and anti-TRDrat (red) at each time point. Fluorescence images for cells were acquired using identical settings and processed offline at the same time, to permit direct comparisons. (A) Representative cells are shown for time points as indicated for anti-TRDdog alone. (B) Immunofluorescence for native TRD (anti- TRDrat) for the16 h time point, typical for all cells. (C) Magnification of outlined box (panel A, 16 h) showing TRDdog appearing first at juxtanuclear sites. Scale bar 2 µm. (D) Immunoblot analysis of cardiomyocyte extracts showing TRDdog and TRDrat with increasing times of TRDdog overexpression, as indicated. The TRDdog immunoblot was counter-stained with anti-TRDrat to show differences in their respective mobilities. Molecular weight markers are shown in kDa on the left.

4.2.3 In situ interaction of JCTdog and TRDdog with CSQ2-DsRed in rough ER

Although newly-synthesized JCT or TRD formed a punctate pattern by 48 h detectable amounts of protein could be observed even closer to the nuclear surface at early time points, suggesting that newly formed junctional SR puncta were distal to the actual sites of biosynthesis.

61

Figure 4-4. TRD and JCT move along a common pathway to populate junctional SR. Cardiomyocytes were not treated (-), or co-treated with TRD and JCT adenoviruses for 18, 24 or 48 h, then fixed for immunostaining and remaining cells harvested for immunoblotting. (A) Adult rat cardiomyocytes coimmuno- stained with anti-TRDdog (green) and anti-JCTdog (red); nuclei are blue. Scale bar, 10 µm. (B) Magnification of outlined box (panel A). Scale bar, 2 m. (C) Upper Stained nitrocellulose transfer of protein extracts from untreated and treated cells at various time points as indicated. Lower Autoradiogram from immunoblot with anti-TRDdog (upper band) and anti-JCTdog (lower band) showing canine protein expression levels. Molecular weight markers are indicated on the left.

To further investigate the sites of junctional SR protein biosynthesis, we compared early JCT and TRD sites of accumulation with the bright red DsRed fluorescence that accompanies accumulation and concentration of CSQ2-DsRed near its site of biosynthesis [85, 171]. Co-expression of either TRD or JCT with CSQ2-

DsRed led to co-localization of their immunofluorescence with CSQ2-DsRed polymers in the nuclear envelop (Figure 4-5 panel A). Prior to formation of CSQ2-DsRed red polymers at 24 h, JCT and TRD showed somewhat typical juxtanuclear patterns of junctional SR filling. However, by 48 h, JCT and TRD had accumulated to high levels in

and around the nuclear envelope, co-localized with CSQ2-DsRed.

62

Figure 4-5. In situ interactions of JCT and TRD with CSQ2-DsRed at juxtanuclear sites. Co-expression of CSQ2-DsRed was carried out with either JCT or TRD for varying times, as indicated, then cells were fixed for immunostaining and remaining cells harvested for immunoblotting. (A) Cardiomyocytes co-expressing CSQ2-DsRed and either JCT, TRD, or delTRD, for 24 or 48 h. Immunofluorescence staining using either anti-JCTdog or anti-TRDdog is shown in green, CSQ2-DsRed is red. Scale bar, 10 mm. (B) Cells were not treated (-) or were treated with JCTdog and CSQ2-DsRed adenoviruses for the indicated number of hours. Immunoblot data for cells were obtained using a mixture of anti- CSQ2 and anti-JCTdog antibodies. (C) Cells were not treated, lane 1; treated with TRDdog and CSQ2-DsRed adenoviruses for 48 h, lane 2; or were treated with delTRD and CSQ2-DsRed adenoviruses for 48 h. Immunoblot data for cells were obtained using a mixture of anti-CSQ2 and anti-TRDdog antibodies. A band that appears in all lanes near 40 kDa (asterisk, *) was not seen in other experiments.

63

Figure 4-6. CSQ2- DsRed-containing ER traffics between myonuclei in line with MTs. Cardio- myocytes were cultured for 48 h with Ad.CSQ2- DsRed adenovirus, and fixed before immunostaining with anti-alpha tubulin (green). (A) CSQ2- DsRed fluorescence shows high levels of accumulation in juxtanuclear ER cisternae, but also within longitudinal tubules of ER extending between the two nuclei. (B) Double labeling of CSQ2-DsRed (red) and α-tubulin immune-fluorescence shows abundant MTs extending between nuclei. Anti-tubulin immunofluorescence (C) of the panel B inset shows a complex scaffold that might support ER tubules containing CSQ2-DsRed (D). Scale bar A, B 10 m and D, 5 m.

To show that newly made TRD was binding to CSQ2 through its well- characterized interaction domain [104], we compared the effect of CSQ2-DsRed on

del trafficking of newly accumulated TRD, a deletion mutant of TRDdog that has absent the canonical CSQ2-binding site. delTRD showed little co-localization or restricted

anterograde movement with CSQ2-DsRed (Figure 4-5 panel A). These data suggested

that CSQ2-DsRed, TRD, and JCT are biosynthesized at a common intracellular site,

likely corresponding to the nuclear envelope. These data also demonstrate that both

JCT and TRD can bind to CSQ2 in situ.

64

Figure 4-7. TRD is retained by its in situ binding to polymerized CSQ2-DsRed in the smooth ER lumen. Cardiomyocytes were cultured with adenoviruses encoding CSQ2-DsRed and TRD for 36- 48 h, which led to co-localized protein accumulation around and between myonuclei. CSQ2-DsRed, once polymerization occurs, serves as a marker for early ER. (A) TRD retained in large part within this series of smooth ER membranes, depending upon the rates of TRD synthesis and trafficking. (B, C) Individual color channels for CSQ2- DsRed (red channel) or TRD (green channel) are shown in black and white (D) At 36 h of expression, CSQ2-DsRed was present in the nuclear envelope, but also present in the ER tubules aligned axially across the cell (white lines in inset. TRD puncta (green) were aligned along these smooth SR tracts suggesting that smooth ER may contribute to sorting of TRD to junctional SR sites (E) Punctate TRD immunofluorescence always co-localized with CSQ2-DsRed in nuclear envelope but some portion generally appeared to outpace CSQ2-DsRed, leading to green puncta with little CSQ2-DsRed fluorescence. Scale bar 10 m, A-C; 5 m, D, E.

4.2.4 Morphological distinctions between smooth ER and junctional SR

In this study, we found that longer periods of CSQ2-DsRed accumulation produced red fluorescent polymers that populated ER tubules visualized along MTs between and surrounding the two myonuclei (Figure 4-6). This CSQ2-DsRed-positive

ER often appeared as circular or polygonal tubules, presumably depending upon its orientation in optical slices. Meanwhile, around nuclei, and particularly between nuclei,

CSQ2-DsRed-containing ER appeared as intertwined tubular and polygonal membranes (Figure 4-7 panels B, D). The underlying MT network in these regions

65 suggested that these more extensive smooth ER components might lie along a dense,

longitudinally directed and looping MT network. (Figure 4-7 panels B, C).

To investigate in more detail the ER accumulation sites of CSQ2-DsRed with those of newly synthesized TRD and JCT, we examined the CSQ2-binding deletion mutant delTRD, to prevent its strong in situ interaction with CSQ2-DsRed. Generally,

after 36 h of incubation with the adenovirus, CSQ2-DsRed had developed its red

polymerization in juxtanuclear cisternae (Figure 4-8 panel A, red fluorescence), during

del which time TRDdog (Figure 4-8 panel A, green fluorescence) has developed a

distribution pattern approaching that of native steady-state junctional SR proteins.

del Expressed alone, TRDdog produced a time-dependent change in fluorescent puncta

that was similar to that of wild type TRDdog and JCTdog proteins (APPENDIX C Figure A-

2). When co-expressed with CSQ2-DsRed however, delTRD immunofluorescence and

DsRed red fluorescence labeled morphologically distinct subcompartments (Figure 4-8

panels B, C). CSQ2-DsRed was localized to the nuclear envelope and smooth

longitudinal ER tubules aligned between and around nuclei along MT-like pathways.

However, delTRD immunofluorescence was generally very low in CSQ2-DsRed-

containing smooth ER, instead concentrating in puncta. Longer incubation times (~72

h) led to increased fluorescence intensity of juxtanuclear tubules of CSQ2-DsRed, yet

junctional SR sites marked by delTRD immunofluorescence still maintained a distinct

separation from DsRed fluorescence.

66

Figure 4-8. The CSQ2- binding mutant delTRD readily crosses the smooth ER to concentrate at sites of junctional SR. Cardiomyocytes were cultured with adenoviruses encoding CSQ2-DsRed and TRD for 48 h. (A) CSQ2- DsRed fluorescence surrounds each myonuclei, but little overlap is seen with green anti-TRD immunofluorescence (delTRD). (B-B2 and C-C2) Enlarged portions of the two perinuclear regions (insets panel A) are shown with the individual channels for CSQ2-DsRed and delTRD. CSQ2-DsRed was highly enriched in areas around and between nuclei, and greatly reduced in more distal ER/SR compartments (panels B1 and C1). TRD immunofluorescence (panels B2 and C2) shows the opposite effect: reduced immunofluorescence in CSQ2-DsRed compartments (smooth ER), with concentrated junctional SR puncta originating in tubules outside of the perimeter of ER defined by CSQ2-DsRed. Transition of smooth ER-to-junctional SR (arrows) (D) TRD after 36 h overexpression can be seen traversing the smooth ER as it traffics toward junctional SR enrichment (E) After 48 h, the smooth ER becomes expanded by CSQ2-DsRed at the same time as junctional SR puncta appear at further distances from the nuclear surface.

Transitions from the more longitudinally oriented CSQ2-DsRed-positive ER, to

del transversely aligned puncta containing TRDdog immunofluorescence, generally

occurred over a radial distance of only a few hundred nanometers. This indicated that

delTRD was retained at lower levels in the proximal CSQ2-DsRed that did not develop into fluorescent puncta. Beyond a defined distance from the central axis of the cell,

67

delTRD began showing punctate junctional SR sites along the MTs, taking on the

punctate appearance of junctional SR (Figure 4-8 panel D). The fundamental difference

between smooth ER-filled tubules and the more distal junctional SR puncta appeared therefore to correspond to whether a deposition and concentration of ER cargo (TRD and JCT) occurred in the vicinity of the Z-line. Since TRD and JCT readily sort and traffic to junctional SR, and polymerized (red) CSQ2-DsRed concentrates almost exclusively in a more proximal ER subcompartment that extends away from juxtanuclear rough ER, we propose that the CSQ2-DsRed is a unique marker of a pre- junctional SR (cardiac rough ER and cardiac smooth ER).

4.2.5 Inhibition of anterograde trafficking by nocodazole

Trafficking of JCT and TRD away from the nuclear envelope occurs within ER tubules that are known to be associated with MTs, along longitudinal and transverse pathways, we predict that trafficking to junctional SR would require a MT platform from which molecular motors could carry the junctional SR cargo.

To determine the effects of MT disruption on early accumulation and initial trafficking steps of JCT and TRD, we used nocodazole. Effects of nocodazole on disruption of junctional SR have been previously reported [188]. Immunofluorescence of both α-tubulin and β-tubulin was examined in the adult rat cardiomyocyte, and differences have been reported in the monomer distribution as shown (Figure 4-9).

Using each of two different anti-tubulin antibodies, a high degree of MT disruption was seen after as little as 2 h incubation with 20 M nocodazole (Appendix C Figure A-3).

68

Figure 4-9. Nocodazole disrupts anterograde movement of junctional SR proteins. Immuno- fluorescence performed on adult rat cardiomyocytes untreated (-) or treated (+) with nocodazole (20 M). Cells were fixed and stained after 48 h in culture. (A) Cardiomyocytes treated with Ad.TRDdog and stained with anti- TRDdog (green). Cardiomyocytes treated with Ad.JCTdog and stained with anti-JCTdog (green). Cardiomyocytes co-treated with Ad.TRDdog and Ad.JCTdog and stained with anti-TRDdog (green) and anti-JCTdog (red) (B) Magnification of outlined box from panel B showing TRDdog. (C) Immunocytochemistry on cardiomyocytes showing α- tubulin (green) and β-tubulin (red). Scale Bar 10 m in A, C, 2 m in B.

Although MT structure was overtly disrupted by 2 h of 20 µM nocodazole, changes in the overall patterns of JCT and TRD puncta were not obvious at the light microscope level (Appendix C Figure A-3). Therefore, to ascertain the effects of MT disruption on JCT and TRD trafficking, we examined whether nocodazole would alter the sites of protein accumulation over the entire period of time during which JCT and

TRD would acquire their normal junctional SR distribution. Nocodazole was therefore added to cells after only 16 h in culture. Such extended periods of MT disruption largely prevented JCT and TRD from leaving their sites of synthesis. These treatments resulted in bright fluorescent rings of juxtanuclear ER clusters as opposed to the anterograde trafficking seen in untreated cardiomyocytes (Figure 4-9 panel B). Levels

69 of JCT (Appendix C Figure A-3 panel C) and TRD (data not shown) remained similar in

nocodazole treated and untreated cells. The data suggested that the disruption was

mostly affecting the transport of newly synthesized proteins and not their steady state distributions.

4.3 Summary

In this chapter, we showed how junctional SR protein biogenesis begins in the nuclear envelope using immunofluorescence analyses of protein expression. Newly synthesized JCT and TRD appeared by 24 h as bright fluorescent puncta close to the

nuclear surface, decreasing in intensity with increasing radial distance. With increasing

time (24-48 h), fluorescent puncta appeared at further distances from the nuclear

surface, resembling steady-state patterns of junctional SR. CSQ2-DsRed prevented

anterograde traffic of newly made TRD and JCT. In situ binding of TRD to CSQ2-

DsRed and effects on its trafficking were abolished if a form of TRD was used in which

CSQ2-binding sites are removed (delTRD).

Double labeling studies confirmed that delTRD first concentrates in junctional SR

puncta that are morphologically distinct from CSQ2-DsRed sites, although delTRD also

accumulated at low levels in these earlier secretory subcompartments. Besides being

localized to the nuclear envelope (rough ER), CSQ2-DsRed labeled a novel smooth ER

network that surrounds nuclei and connects the nuclear axes.

The transmembrane junctional SR proteins transit across perinuclear smooth ER

that extend along microtubules away from the nucleus, concentrating only where ER

trafficking crosses Z-lines. ER/SR protein traffic required MTs, as nocodazole treatment

led to disruption and minimal anterograde protein movement of both JCT and TRD, with

accumulations close to the nuclear envelope.

70

CHAPTER 5

TRAFFICKING OF THE LUMINAL SR PROTEIN CSQ2

5.1 Rationale

The luminal junctional SR protein CSQ2 was discovered many years prior to the

transmembrane CSQ2-binding proteins JCT and TRD (Chapter 4), and in many ways is

the most enigmatic of the group of proteins classically known to enrich in junctional SR

subcompartments. As a luminal protein, it is not restricted by the dynamics of the lipid

bilayer, and should be capable of faster trafficking through secretory pathway lumens.

At the same time, because it contains no known retrieval sequences, it should in principle be secreted from the cardiomyocyte - but apparently is not. We have shown

that the retention of different forms of the protein CSQ (e.g. CSQ1, CSQ2, and CSQ2-

DsRed) is uniquely determined by their polymerization, which is sensitive to levels of

Ca2+ and ionic milieu [19].

ER trafficking of luminal cargo requires communication with cytosolic trafficking proteins. To define the biosynthetic trafficking pathway for luminal junctional SR

proteins, we expressed canine CSQ2 and detected it with canine-specific protein

antibodies. These studies extend the data obtained for JCT and TRD, and present a

wider view of junctional SR protein trafficking and junctional SR compartmentation.

Hypothesis: CSQ2, a luminal junctional SR protein is synthesized in the juxtanuclear

rough ER and traffics through the same ER/SR pathways defined by our studies on the

transmembrane proteins JCT and TRD.

5.2 Results

5.2.1 Detection of CSQ2dog using species-specific antibodies

To determine the specificity of the antibody utilized in the detection of expressed

71 canine CSQ2, immunoblot analysis was performed on protein samples from canine and rat microsomal vesicles. In comparison to canine microsomes where CSQ2 was detected, no band was detected in rat microsomes using anti-CSQ2dog antibody (Figure

5-1 panel B). Thus, this data shows that the antibody used for detection of CSQ2 is highly specific to the expressed canine form with relatively no cross reactivity with the rat form of the protein. Meanwhile, rat forms of CSQ2 can be detected using a second antibody that reacts with both canine and rat forms.

When immunocytochemistry was performed on adult rat cardiomyocytes expressing CSQ2 for 48 h, CSQ2dog co-localized with the endogenous native rat form of the protein at junctional SR sites and displayed a similar subcellular distribution (Figure

5-1 panel C). However, displayed a similar but antibody specific staining.

Figure 5-1. Specificity of anti-CSQ2 antibodies. Samples (50 g) of canine heart microsomes (MVc) and rat heart microsomes (MVr) were analyzed by SDS-PAGE (10% acrylamide) and transferred to nitrocellulose. (A) Amido black stained nitrocellulose. (B) Immuno- blotting of CSQ2 (arrowhead) using anti-CSQ2dog and anti-CSQ2rat antibodies. (C) Immunofluorescence C of CSQ2 in cultured adult rat cardiomyocytes treated with Ad.CSQ2 for 48 h using anti-CSQ2 (green) and anti-CSQ2rat (red). Scale Bar 10 m.

5.2.2 Determination of CSQ2 initial sites of accumulation

To determine the biosynthetic pathway luminal junctional SR proteins follow, a time course of canine CSQ2 expression was performed over 48 h in adult rat

72 cardiomyocytes. To visualize the CSQ2 accumulation, cultured rat adult cardiomyocytes were analyzed after 16, 24, 30 and 48h of overexpression. Confocal images of indirect immunofluorescence were generally acquired as a Z-stack covering the height of the cell in contiguous series of optical slices of 0.5 m thickness.

In contrast to accumulation at juxtanuclear sites for the transmembrane junctional

SR proteins TRD and JCT, CSQ2 was widely distributed throughout the cell after only

16 h, accumulating at junctional SR sites with no concentration around the nucleus

(Figure 5-2). We believe that the rapid transit and approach to steady state distribution reflects the differences between the transport of luminal and transmembrane ER/SR proteins. Thus far, these data do not support our hypothesis that all junctional SR proteins are made in a common juxtanuclear sites

Also, unlike the distinct juxtanuclear localization of the fusion protein CSQ2-

DsRed (see Chapter 4 section 4.2.2), CSQ2 (wild-type) was widely distributed. The differences in localization for wild-type versus CSQ2-DsRed likely reflect their polymerization-dependent localizations. This would explain why CSQ2-DsRed remains in rough ER while native CSQ2 traffics more distally to contiguous junctional SR sites.

Given the rapid transit of CSQ2 across ER /SR compartments, we hypothesized that the luminal protein trafficking was too rapid to permit juxtanuclear accumulation.

Therefore, to visualize the earliest sites of CSQ2 accumulation, overexpression of

CSQ2 was performed at higher adenoviral concentrations 2x and 5x.

73

Figure 5-2. Time course of CSQ2 accumulation at junctional SR sites. Coverslips containing cardiomyocytes were removed and fixed following treatment with Ad.CSQ2 for 16, 24, 30 and 48 h and remaining cells harvested for immunoblotting. (A-D) Immunofluorescence of CSQ2 (green) and nuclei (blue) at different time points, cells were stained with anti-CSQ2dog. Scale bar 10µm. (E) Amido Black stained nitrocellulose of protein extracts from cardiomyocytes corresponding to 16, 24, 30 and 48 h of Ad.CSQ2 treatment. (F) Immunoblot analysis of cardiomyocyte protein extracts using anti-CSQ2 showing CSQ2 levels with increasing times of canine CSQ2 overexpression. Molecular weight markers (kDa) indicated on the left.

In contrast to CSQ2 1x adenoviral treatments, higher juxtanuclear CSQ2 immunofluorescence signals were detected in cardiomyocytes that received 2x and 5x

Ad.CSQ2 (Figure 5-3). The distribution of CSQ2 fluorescence at 5x Ad.CSQ2 treatments displayed a membrane-subcompartment-filled in juxtanuclear membrane sites and not a strictly punctate pattern as seen at distal sites. These data support the idea that CSQ2 is also synthesized in and around the nuclear envelope, although the levels of overexpression needed to visualize it are too high to warrant detailed analysis.

Interestingly, the changes in ER morphology that accompanied the highest levels of

CSQ2 overexpression in the juxtanuclear area appear somewhat similar to the increased ER cisterna observed with CSQ2-DsRed expression (Chapter 4 section

4.2.4).

74

Figure 5-3. Higher rates of CSQ2 biosynthesis create higher fluorescence intensities at juxtanuclear sites. Coverslips containing cardiomyocytes were removed and fixed following treatment with Ad.CSQ2 for 48 h at 1x, 2x and 5x, remaining cells were harvested for immunoblotting (A) Immunofluorescence of CSQ2 (green) in adult rat cardiomyocytes detected using anti-canine CSQ2, following 1x, 2x and 5x Ad.CSQ2 treatments for 48 h. Nuclei (blue) are stained with DAPI. Scale bar 10m (B) Enlarged portions of the perinuclear regions (panel A, insets) (C) Autoradiogram showing CSQ2 levels. Protein extracts are from cardiomyocytes cultured for 48 h either untreated or treated with Ad.CSQ2-WT at 2x and 5x, respectively. Molecular weight markers in kDa indicated on the left. (D) Fluorescence intensity of CSQ2. Quantification of fluorescence from CSQ2dog expression (in 10 cells /condition) was measured as a function of transverse distance (m) from the nuclear edge in 48 h cultured cardiomyocytes. Values of fluorescence intensity are relative to the intensity at the edge of the cell.

5.2.3 In situ interaction of TRDdog with CSQ2

Previous studies have demonstrated in vitro interaction between proteins that are

75 part of the Ca2+ release complex. Moreover, we have shown here that CSQ2-DsRed

expression led to the complete retention of TRD and JCT in and around the nuclear

envelope. We therefore, sought to determine whether CSQ2 accumulation, localization

and rates of trafficking would be affected by the overexpression of TRD or vice versa.

Primary adult rat cardiomyocytes were co-treated with Ad encoding canine CSQ2 and

TRD. Expressed protein levels were detected using canine specific anti-CSQ2 and anti-

TRD, immunoblotting showed an increase in protein levels at 48 h compared to 24 h

(Figure 5-4, panel A). Immunocytochemistry showed no accumulation of CSQ2 at

juxtanuclear sites upon co-overexpression with TRD (Figure 5-4, panels B, C), unlike

CSQ2-DsRed co-overexpression that concentrated TRD at juxtanuclear rough ER sites.

The slower trafficking transmembrane TRD did not exhibit an overt effect on the faster

trafficking of luminal CSQ2. This may be due to the rapid passage and dispersal of

CSQ2 through the SR/ER lumen away from the membrane surface and the nuclear

envelope.

5.2.4 Effect of nocodazole on junctional SR sites and CSQ2 trafficking

MTs are known to interact with the ER and are involved in the extension of ER tubules. To determine the effect of nocodazole on CSQ2 accumulation at junctional SR sites, cells expressing both CSQ2dog and TRDdog were treated with 20 M nocodazole

(Figure 5-4, panel D). Immunofluorescence analysis showed a distinct effect on the

transmembrane protein TRDdog but with minimal to no effect on the expressed luminal protein CSQ2. Nocodazole also had no effect on endogenous junctional SR containing

CSQ2 localization upon nocodazole treatment (Appendix C Figure A-4). The lack of effect of nocodazole on CSQ2 trafficking or native junctional SR puncta compared to

TRD anterograde trafficking may be due to the presence of multiple cytoskeletal

76 structures that help to maintain the more distal subcellular membrane compartments

[127].

Figure 5-4. CSQ2 maintained independent transit when co-expressed with TRD and was not similarly disrupted by nocodazole treatment. Coverslips containing cardiomyocytes were removed and fixed following culture for 24 or 48 h and co-treated with both Ad.CSQ2, Ad.TRD and with or without nocodazole, remaining cells were harvested for immunoblotting (A) Autoradiogram showing protein levels of expressed CSQ2 and TRD. Protein extracts were obtained from cardiomyocytes co-treated with Ad.CSQ2 and Ad.TRD for 24 and 48 h. Proteins detected using anti-CSQ2dog, and anti- TRDdog. Molecular weight markers (kDa) indicated on the left. (B-D) Immunofluorescence of CSQ2 (green) and TRD (red) in cardiomyocytes co-treated with Ad.CSQ2 and Ad.TRD for 24 h or 48 h. (B) Cultured for 24 h, (C) Cultured for 48 h (D) cultured for 48 h and treated with nocodazole (20 µM). Nuclei (blue) are counter-stained with DAPI. Scale bar 10 m.

5.3 Summary

In this study, we examined the luminal SR protein CSQ2 biogenesis. CSQ2 established steady state distribution and did not show juxtanuclear accumulation at 1x

CSQ2dog expression. Sites of biosynthesis were less prominent for CSQ2 because it is

77 luminal (compared to TRD or JCT), and the wild-type forms of CSQ2 cannot polymerize in the juxtanuclear compartment. Nevertheless, higher CSQ2 expression levels were able to validate the hypothesis that junctional SR proteins have a common biosynthetic/secretory pathway. The data provides a consistent model of ER and SR

transmembrane (Chapter 4) and luminal (Chapter 5) protein transport across the

cardiomyocytes, and a new understanding of pathways by which cellular Ca2+ handling

protein complexes can be maintained and modified.

78

CHAPTER 6

DISCUSSION

Crucial distribution and localization of proteins within the adult cardiomyocyte inner membrane system are the result of the ongoing sorting and trafficking of proteins.

Many resident proteins of the ER/SR and sarcolemmal membranes continually take part in this membrane protein traffic, yet very little is known about these processes in the heart. A range of biochemical and immunological techniques were used in this

Dissertation to develop a basic model of protein trafficking in the cardiomyocyte that may also explain fundamental changes observed in intact hearts during failure.

6.1 CSQ2 mass redistribution in two types of cardiomyopathies

In Chapter 3, we evaluated CSQ2 purified from heart tissue following two different types of induced cardiomyopathies. Remarkably, we found similar patterns of mass redistribution in the two disparate types of cardiomyopathies when compared to C mongrel dogs. The changes in mongrel dogs mimicked the changes previously reported for tachypaced beagles [178], with a series of interesting new findings.

Overall, our results suggest that a common mechanism for change in CSQ2 mass

(glycosylation and phosphorylation) and turnover exists that is independent of the

pathologies that lead to the syndrome of HF (Figure 3-1 panel b), likely reflecting an

adaptive response of the distressed myocardium. These studies involved changes in

two cardiac pathological mechanisms by which an animal is put into heart failure,

apparently manifested through a common biochemical and cell biological change.

Hearts from HFi dogs reportedly exhibit increased LV wall thickening (27%) and

increased heart weight to body weight ratios (18%) [165], whereas tachypacing is

marked by LV wall thinning, LV dilatation, and reduced heart weight [184]. HFt hearts

79 can develop LV hypertrophy following resumption of normal pacing [189], suggesting

that a single program of remodeling develops that is independent of the induction route.

On the other hand, the different types of hypertrophy reported for HFt and HFi tissues

might explain the similar but distinct changes in CSQ2 mannose trimminig reported

here.

The single biological change might be related to changes known to occur as part

of cardiac hypertrophy. Mechanisms for cardiac hypertrophy remain unclear but are

likely to involve formation of new Ca2+ microdomains within the ER or SR of the inner

membranes of the cardiomyocyte [112, 190]. Changes in CSQ2 glycan structure in

failed heart tissue correspond to changes brought about by the formation of new protein

that undergoes less glycan processing.

CSQ2 molecules in the canine heart undergo co-translational N-linked

glycosylation on 316Asn [167]. In addition, canine CSQ2 molecules are phosphorylated

on 0,1,2, or 3 serines contained within the CSQ2-specific C-terminal cluster 378,382,386Ser

[167, 191]. This modification is thought to be carried out by a cytosolic protein kinase

CK2, also occurring co-translationally [183]. Therefore, the greatest posttranslational

mass for CSQ2 is 47,478 Da, which contains 9 mannoses and 3 phosphates. This form

of CSQ2 ordinarily only accumulates at high relative levels when canine cDNA is adenovirally-overexpressed in non muscle cell lines [167]. In contrast, very little of this

unprocessed form is found in the mammalian heart [167, 179]. Most CSQ2 in the heart

is highly mannose-trimmed and relatively dephosphorylated [179]. We believe that he

degree of posttranslational processing is likely to reflect critical regulatory roles in CSQ2

turnover. Mannose trimming of N-linked glycans is likely to reflect CSQ2 degradation

[180, 181], whereas CK2 phosphorylation states may reflect CSQ2 turnover [179, 183].

80

The mechanisms underlying the CSQ2 mass redistribution are likely to lie in

CSQ2 cellular processing, from its biosynthesis, to its accumulation in ER/SR

subcompartments, to its degradation. One possibility for the changes is that the two

accumulated pools emanate from two distinct sites of CSQ2 biosynthesis (rough ER),

one that is roughly common to both normal and HF myocardium (Pool I), and one that is

activated only in cardiomyopathy states of the myocardium (Pool II). The lack of

posttranslational processing of Pool II suggests either that this CSQ2 is not being

transported to junctional SR, or perhaps degrades before undergoing further mannose

trimming. In studies aimed at elucidating sites for junctional SR protein synthesis

(Chapter 4 and 5), we found no evidence to support more than a single subcellular site.

We cannot however, exclude the possibility that additional sites of biosynthesis arise or

become activated in cardiomyopathy.

An immunofluorescence analyses of thin tissue sections was aimed at

determining whether quantifiable differences could be detected across the cell, from

juxtanuclear rough ER transversely to the cell surface. The lack of significant changes

in CSQ2 immunoreactivity in C and HF tissue may suggest that a more complex

subcellular distribution underlies the observed mass redistribution. In our new

trafficking model, CSQ2 and other SR proteins could exhibit differences in a smooth ER

to junctional SR step that could also explain the changes in its glycan structure. It will

be interesting to extend the immunofluorescence analyses to include changes that are

reflective of MT trafficking or smooth ER transport; new ideas that based upon our

proposed model of membrane protein trafficking.

The two CSQ2 mass pools could also develop from a single site of CSQ2

biosynthesis, after which differences occur in the rates of anterograde trafficking.

81

Different rates of anterograde transit through the cardiac secretory pathway might result from different rates of CSQ2 translocation through CK2-modified translocons [183, 192,

193], or as a result of increased retention of CSQ2 transcripts in rough ER following

CK2 phosphorylation [183]. We previously reported that CSQ2 in the failing beagle heart showed increased phosphorylation on protein kinase CK2-sensitive sites [178], and we have also shown that CK2 phosphorylation of CSQ2 can result in increased ER retention [183]. Calnexin, another resident ER/SR protein substrate for CK2, may associate with the translocon upon CK2 phosphorylation [193]. A broader role for CK2

activation of cardiac hypertrophy was recently predicted, based upon a hypertrophic

activation of p27/Kip1, which leads to CK2 dissociation and activation [194]. This model

of cardiac hypertrophy involves the intriguing connection between p27/Kip1 activation of

cell proliferation (GS-phase transition) in dividing cells, and activation of hypertrophy

in the non-dividing cardiomyocyte.

Finally, accumulation of CSQ2 in a second metabolic pool might represent the

effects of T-tubule changes. Loss of normal T-tubule morphology has been widely

reported in HF based upon microscopic visualization in tissue and cultured cells of mammalian HF models [195-197]. Retraction of the transverse invaginations of

sarcolemma leaves a disorganized ultrastructure with orphaned RyRs that contribute to arrhythmogenesis through under-regulated sparks [195]. With loss of triggered Ca2+ release, CSQ2 might play a more prominent role in regulation of SR Ca2+ release in

RYRs orphaned by T-tubular reorganization. Junctophilin-2, a junctional SR protein that

forms a critical connection to T-tubules, is thought to play a role in T-tubule

reorganization [198]. How and whether such changes would affect other junctional SR

proteins or CSQ2 processing is not known. Data reported here would represent the first

82 evidence for specific changes in junctional SR protein processing with T-tubular change

in vivo.

6.2 Junctional SR subdomains: protein trafficking and accumulation

In the studies described in Chapter 4 and 5, we overexpressed JCT, TRD and

CSQ2 in cultured adult rat cardiomyocytes. A putative transport pathway that would

populate successive junctional SR subcompartments has not been previously

described, primarily because little attention has been given to the specialized cell

biology of the cardiomyocyte. In these studies, we were able to show evidence for

synthesis in and around the nuclear envelope. In addition, we established CSQ2-

DsRed as a marker for a novel compartment based upon double labeling with either

TRD or delTRD.

Studies of SR subcompartment biogenesis have mostly examined the change in

subcompartmentation during muscle cell differentiation, and while those studies can

provide insights, they are not aimed at the fundamental questions of cardiac SR biology

of the adult cardiomyocyte. In our studies we specifically studied primary adult

cardiomyocytes. Newly synthesized TRD and JCT generated immunofluorescent

puncta close to the nuclear surface 18-24 h post-adenoviral expression, as did CSQ2

but only at accelerated rates of synthesis. Data obtained for TRD, JCT and CSQ2

corroborate previous findings of CSQ2-DsRed synthesis within cisternae that surround

myonuclei, generating a model of common trafficking for all junctional SR proteins.

In situ interactions between TRD or JCT and CSQ2-DsRed, and effects of

nocodazole on trafficking, suggested that a smooth ER likely underlies all membrane

protein trafficking radially and axially away from the nuclear envelope, maintaining

junctional SR protein levels through selective retention. ER transmembrane protein

83 traffic requires intact MTs, as nocodazole treatment led to loss of anterograde JCT and

TRD traffic away from the nuclear envelope. Disassembly of MTs prevented

anterograde trafficking, leading to build-up of protein in the nuclear envelope as

predicted by our general model.

Given that CSQ2-DsRed maintains ability for rapid in situ Ca2+-binding and release around the nucleus [171], we reasoned that the fusion protein CSQ2-DsRed might affect its well-established in vitro binding partners TRD and JCT. Indeed, rough

ER-localized CSQ-DsRed was able to form a veritable net that prevented anterograde trafficking of JCT and TRD. Upon co-expression of CSQ2-DsRed with TRD and/or JCT, anterograde trafficking to junctional SR was completely inhibited, and JCT and TRD were retained within the nuclear envelope and juxtanuclear membranes. Under these

conditions, JCT and TRD did not show the punctate staining of junctional SR, but

collected to a high level in juxtanuclear cisternae characteristic of CSQ2-DsRed and the

del rough ER. Lack of CSQ2-DsRed binding to TRDdog, prevented retention of TRD in the

nuclear envelope, but did not alter its concentration in junctional SR puncta, suggesting

that the KEKE-motif that binds CSQ2 in vitro [104] in TRD (and presumably JCT) is not

necessary for its targeting and concentration in junctional SR. Wild-type CSQ2,

however, was not retained by the slower-trafficking TRD. Since in situ interactions were

shown to be able to occur, the lack of interaction between newly made transmembrane

TRD and the newly made luminal CSQ show how protein interactions within the

secretory pathway require some spatial coordination, a situation that must occur in vivo.

The amount of time needed for TRD and JCT to accumulate to levels sufficient

for confocal imaging (12-16 h), and the time intervals of ~12 h needed for seeing

changes in protein localization indicative of its trafficking, reflect the passage through

84

the lipid bilayer and across longitudinal tubules of ER. JCTdog and TRDdog immunofluorescence traverses the nuclear envelope (rough ER) with little retention, yet areas of concentrated protein can be seen in the nuclear envelope, which appear to represent an initial sorting step from which transit is launched. Visualization of JCTdog or TRDdog within the nuclear envelope required binding it to the CSQ2-DsRed polymer

that never leaves the ER, or addition of nocodazole to disassemble its anterograde escape route.

An alternative explanation for the observed time-dependence in accumulation is a rate-limiting mRNA transport step. In such a model of our data, RNA would have been transcribed and spliced close to the nuclear surface; mRNA would then be transported along MTs aligned along Z-lines. In this paradigm, protein synthesis would presumably occur at successive junctional SR puncta over 48 h. Metsikko and co- workers hypothesized a specialized mechanism for “Ca2+-handling” proteins in cultured

skeletal muscle myotubes. They described a subcellular localization for CSQ1 and

SERCA mRNA using electron microscopic in situ hybridization [199]. However, the

del differential binding of TRDdog and TRDdog to perinuclear CSQ2-DsRed, indicate that

the rate of TRD accumulation across the cell interior is dependent upon anterograde

protein traffic instead of a putative cytosolic mRNA transport.

Smooth extensions of the rough ER membrane traffic proteins away from the

nucleus along microtubules radially along Z-lines, as well as longitudinally (axially). The data suggest that retention of junctional SR cargo proteins near T-tubules occurs via this ER traffic. ER tubule extension may occur in response to MT polymerization through attachment of ER to MT tips. Membrane trafficking and organization has been linked to microtubule organization [146, 200]. MTs and the molecular motors dynein

85 and kinesin are thought to regulate the dynamic changes that are part of steady-state

SR biology [188]. The most common mechanism for MT-based transport is through cargo association either directly or indirectly with motor proteins [201]. We hypothesize that following protein maturation in juxtanuclear rough ER cisternae, ER tubules or vesicles containing transmembrane JCT and/or TRD traffic towards Z-lines along MTs

(Figure 4-9). Here we showed that upon 36 h treatment with the MT disrupting agent nocodazole, newly synthesized TRD and JCT failed to traffic beyond the nuclear envelope, so that newly made JCT and TRD failed to develop and populate characteristic junctional SR puncta. This disruption of junctional SR population suggests that a critical MT motor-associated transport is essential for delivery of these newly synthesized proteins to the junctional SR sites.

Vega et al examined motile SR boutons in the SR of adult ventricular myocytes labeled by Ad overexpression of red fluorescent protein fused to the well-known ER- retrieval tetrapeptide (RFP-KDEL) [188] (-KDEL ER retrieval discussed in Chapter 1, section 1.4.4). Their data show that RFP-KDEL fluorescence labels the nuclear envelope to high levels, and along transverse membranes. Live cell imaging of motile

(corbular) SR boutons in the cell periphery that contain RyR showed longitudinal movements across Z-line boundaries over the course of several minutes, providing evidence of longitudinal trafficking once junctional SR proteins have populated sites in the cell periphery. Although longitudinal trafficking was not explicitly measured in our experimental system, such protein movements were inferred from the double labeling studies carried out with CSQ2-DsRed and TRD/delTRD. Because the ER exit site

marker sec23 is enriched in a similar Z-line pattern in these cells [85] we previously

suggested that longitudinal traffic away form Z-lines might represent a cardiac ERGIC.

86

The current studies offer important additional insights into this question. We showed here that CSQ2-DsRed remains associated with ER tubules that remain distinct from

junctional SR for at least the 48 h that we recorded its location (this distinction extended

to at least 72 h, N. Sleiman and S. Cala, unpublished observations). The mechanism of

smooth ER to junctional SR transition (Chapter 4 Figure 4-8) remains unknown, but

would be consistent with a role of ER exit sites and the reported build-up of sec23

immunofluorescence along Z-lines [85].

Protein synthesis is a prerequisite for cardiomyocyte health, but accelerated

protein synthesis is a marker of cardiac hypertrophic growth. In cardiomyocytes, the SR

plays an important role in contractile Ca2+ handling compared to other cell types, and

therefore is more extensive and complex. In Chapter 4, we observed an apparent

proliferation of ER following CSQ2-DsRed overexpression. These juxtanuclear

junctional SR sites might play a role in the maintenance of cardiac hypertrophy. The

distance (roughly 1 m) from the nuclear edge is theoretically close enough to serve as

the Ca2+ microdomain hypothesized to support nuclear transcription associated with the hypertrophic state [66, 190]. In support of this idea, we previously reported that the

Ca2+ that is bound to CSQ2-DsRed in adult cardiomyocytes is sufficient to elicit

localized perinuclear Ca2+ transients, and in neonatal cardiomyocytes is sufficient to

cause hypertrophic growth determined as increased surface area [171]. Thus, these juxtanuclear rough ER cisternae and tubules might play a role as a localized Ca2+ microdomain to support Ca2+-dependent transcription thought to drive cardiomyopathic forms of hypertrophy.

6.3 Model of the inner membrane system in the adult cardiomyocyte

Based upon our extensive fluorescence microscopy data using the unique ER

87 and SR markers developed in our studies, we propose an overall model of protein trafficking to junctional SR. The protein markers reflect a series of inner membrane subcompartments and their cell biological relationships. The most important elements of

our proposed model are as follows:

1. Junctional SR proteins are synthesized in the nuclear envelope, generally in both

nuclei.

2. Rough ER is connected to an extensive and plastic smooth ER that connects the

two perinuclear regions. This smooth ER is comprised of polygonal tubules

between the nuclei, but also includes extensive tubules or cisternae that align

longitudinally.

3. Population of junctional SR subcompartments arises from longitudinally aligned

smooth ER that circumvents the cell but is also prominently filled via transversely-

aligned membranes.

4. Smooth ER appears to expand with increasing levels of CSQ2-DsRed expression,

and in so doing, shifts junctional SR subcompartments to more peripheral areas of

the cell.

Proteins destined for inclusion into membrane structures are synthesized on

membrane bound ribosomes (rough ER). Critical processes known to occur in rough

ER include signal sequence docking, mRNA translation, protein translocation into the

ER, N-linked glycosylation, and folding. In the adult cardiomyocyte, the predominant

site of junctional SR protein biosynthesis occurs in close proximity to the nucleus,

presumably within the nuclear envelope. Smooth ER can be comprised of tubules and

flattened cisternae that are free of ribosomes.

Sorting of newly-synthesized proteins to their final destinations requires elaborate

88 specializations of the ER such as transitional ER and Golgi. Transitional ER is a major

ER subcompartment, free of ribosomes. Along smooth ER, transitional ER forms sites

of ER exit, from where an ERGIC can form comprised of fused ER-Golgi transport

vesicles and tubules. In cardiomyocytes, the cytoskeletal scaffold is involved in the

proper localization of proteins (e.g. ion channels) to specific domains of the

sarcolemma. The proper expression of ion channels in the heart is charted by

cytoskeletal components specifically, MTs. Not much is known of how these cargo are

elected and targeted. The extent of MTs involvement and disruption in the localization

or delivery of transmembrane junctional SR was dramatic. To improve the

understanding of cardiac protein synthesis and trafficking and thus the remodeling that

accompanies cardiomyopathies that we consider involves this subcellular remodeling of

intracellular membranes.

Figure 6-1. Schematic model of junctional SR filling by newly-made junctional SR proteins. In adult cardiomyocytes (Normal), protein translation into rough ER membranes occurs in or near the nuclear envelope (NE). Areas exist within the NE (green ellipses) in which translocon complexes undergo alignment with sarcomeres to transport jSR proteins radially (vertical blue arrows). Rough ER becomes sheets or tubules of smooth ER (curved and longitudinal turquoise arrows) that, along with Z- tubules, promote an axial filling of jSR puncta (pink circles). A convoluted set of ER tubules occupy an axis between nuclei (*asterisk). Following high levels of CSQ2- DsRed expression and smooth ER accumulation, smooth ER no longer effectively transitions to jSR sites, leading to a changes in ER and SR membrane partitioning (gray fill).

89

6.4 Conclusions

In cardiomyopathies capable of producing HF, some change in the pathway for

CSQ2 occurs. It can be detected by changes in CSQ2 processing, and its regulation

(perhaps by CK2 phosphorylation in rough ER) may regulate a relative transition of ER form its contractile role to a more transcriptional and hypertrophic function. Whether mass redistribution corresponds to SR morphological remodeling, or is regulated at the level of CSQ2 turnover will be interesting questions for future investigations. It will be important to establish which of the several possible mechanisms underlie the altered processing of CSQ2 in the myocardium, as these in vivo data may add significant insights into adaptive changes in junctional SR physiology and Ca2+ homeostasis in the

heart.

Changes in TRD and JCT accumulation occurring in cardiomyopathies show the

value of understanding the protein trafficking pathways accompanying the biosynthesis

of functional SR membranes. This study demonstrates a novel biosynthetic pathway in

adult cardiomyocytes. We show that junctional SR Ca2+-release complex protein

components arrive at junctional SR proximately following synthesis at juxtanuclear

rough ER sites. Disruption of the normal trafficking cycle along with altered cellular

localization could account, in part, for select cardiomyopathies and changes in Ca2+ homeostasis.

Knowledge of pathways leading to functional ER subcompartmentation will provide a basis for identifying new mechanisms by which protein glycosylation and phosphorylation, and thus trafficking, lead to changes in functional Ca2+ release sites.

Future studies should focus on understanding the regulatory mechanisms in cardiac

membrane trafficking changes that occur in disease states, which may be either

90 adaptive or maladaptive.

91

APPENDIX A

Springer License Terms And Conditions

92

APPENDIX B

Approved Animal Protocol

93

APPENDIX C

Supplemental Data

Figure A-1. Levels of endogenous rat CSQ2 and JCT proteins at various time points in culture. Primary adult rat cardiomyocytes were harvested after 16, 24, 36 or 48 h (A) Amido Black stained nitrocellulose membrane of protein extracts from cultured cardiomyocytes treated with Ad.JCT (+) or non-treated (-) run on Bis-Tris MIDI gel (10 %) (B) Immunoblot of CSQ2rat using anti-CSQ2rat (C) Immunoblot of JCT detected using anti-JCTrat.

Figure A-2. Time course of del TRDdog accumulation at junctional SR sites. Coverslips containing cardiomyocytes were removed and fixed following del treatment with Ad. TRDdog for 18 and 36 h and remaining cells harvested for immunoblotting. (A-C) Immunofluorescence TRDdog (red) and nuclei (blue) at different time points, cells were stained with anti-TRDdog. Scale bar 10µm.

94

Figure A-3. The effect of nocodazole concentration and duration of treatment. Immunofluorescence performed on adult rat cardiomyocytes treated with nocodazole as indicated, fixed and stained with anti-JCTdog (red) after 48 h in culture. (A) Cardiomyocyte expressing JCTdog and treated with nocodazole for 2 h (B) Cardiomyocyte expressing JCTdog and treated with nocodazole for 12 h (C) Autoradiograms of immunoblot performed on protein extracts from cardiomyocytes treated with Ad.JCT for 48 h either with or without 20 M nocodazole for 36 h showing JCTdog and α-tubulin levels.

95

Figure A-4. Junctional SR sites distribution depicted by CSQ2rat endogenous protein localization upon nocodazole treatment. Cultured cardiomyocytes co-stained with anti- CSQ2rat (red) and anti-α-tubulin (green). (A) Cardiomyocyte without nocodazole treatment (A1) α-tubulin, green channel from panel A (A2) red channel from panel A showing CSQ2rat (B) Cardiomyocyte treated in culture with nocodazole (20 M) (B1) α-tubulin, green channel from panel B (B2) red channel from panel B showing CSQ2rat.

96

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ABSTRACT

JUNCTIONAL SARCOPLASMIC RETICULUM PROTEIN PROCESSING AND TRAFFICKING IN CARDIAC TISSUE AND PRIMARY CULTURED CARDIOMYOCYTES

by

NA’AMA H. SLEIMAN

August 2014

Advisor: Steven Cala Ph.D.

Major: Physiology

Degree: Doctor of Philosophy

Junctional SR is an important and unique ER subdomain in the adult myocyte that releases Ca2+ through the actions of an exclusive set of resident proteins. Cardiac calsequestrin (CSQ2) undergoes two co-translational modifications: N-linked glycosylation on 316Asn, and phosphorylation by protein kinase CK2 on a cluster of 3 serines in its tail. In the heart, CSQ2 molecules undergo extensive mannose trimming by ER mannosidase(s), a posttranslational process that often regulates protein breakdown. To investigate CSQ2 protein processing in cardiomyopathy models, studies were performed to test whether CSQ2 glycan structures would be altered in heart tissue from mongrel dogs induced into heart failure (HF) by two different experimental treatments, rapid ventricular pacing or repeated coronary microembolizations. CSQ2 mRNA and protein levels were determined, and its processing was analyzed by electrospray mass spectrometry. Despite the different cardiomyopathies producing the failure, roughly one-third of CSQ2 molecules from each of the failed tissues contained glycan structures indicating a buildup of newly made protein with highly reduced glycan trimming. Analyses of tissue samples showed

120 decreases in CSQ2 protein levels per unit levels of mRNA for tachypaced heart tissue, also indicative of altered turnover.

To place these data in context of junctional SR cell biology, we carried out a series of studies on primary rat cardiomyocytes to understand the basic features of junctional SR protein trafficking. We expressed canine forms of junctin (JCT), triadin

(TRD) and CSQ2 in adult rat cardiomyocytes, each a major protein component of the

Ca2+ release complex. Two additional reagents employed were CSQ2-DsRed, which fails to leave sites of early synthesis due to immediate polymerization, and so was able to bind TRD and JCT in rough ER; and a deletion mutant of TRD (delTRD) which was not so affected, and allowed us to directly probe the differences between ER and SR by double labeling. A complex picture emerged, in which all junctional SR proteins are synthesized in the nuclear envelope but travel across MT linked smooth ER to continuously deposit junctional SR proteins. The data suggest that ER proteins traffic radially along Z-lines, and axially between nuclei, maintaining junctional SR protein levels through sorting and selective retention as ER traverses Z-lines. ER protein traffic requires intact MTs, as nocodazole treatment led to loss of anterograde JCT and TRD traffic away from the nuclear envelope.

We conclude that altered processing of CSQ2 may be an adaptive response to the myocardium under stresses that are capable of inducing heart failure. We provide a model of ER and SR protein transport across the cardiomyocytes, and a new understanding of pathways by which cellular Ca2+ handling protein complexes can be maintained and modified.

121

AUTOBIOGRAPHICAL STATEMENT

NA’AMA H. SLEIMAN

Education Doctor of Philosophy, Physiology, Wayne State University, Detroit, Michigan (August, 2014) Master of Science, Molecular Biology, Lebanese American University, Beirut, Lebanon Bachelor of Science, Biology, American University of Beirut, Beirut, Lebanon

Honors and Awards Thomas C. Rumble University Graduate Fellowship – Wayne State University Education Committee Travel Award - Biophysical Society Graduate Student Professional Travel Award-Wayne State University Physiology Department Graduate Student Representative-Wayne State University Interdisciplinary Biomedical Sciences Fellowship-Wayne State University

Peer Reviewed Publications 1. Jacob S*, Sleiman NH*, Kern S, Jones LR, Sala-Mercado JA, McFarland TP, et al. (*co-first authors) Altered calsequestrin glycan processing is common to diverse models of canine heart failure. Mol Cell Biochem. 2013;377:11-21. 2. McFarland TP, Sleiman NH, Yaeger DB, Cala SE. The cytosolic protein kinase CK2 phosphorylates cardiac calsequestrin in intact cells. Mol Cell Biochem. 2011;353:81-91.

Abstracts 1. Microtubule integrity is essential to junctional SR protein delivery. Sleiman, N.H, Smoczer, C., Cala, S.E. 58th Annual Meeting of the Biophysical Society, San Francisco, CA, 2014, 14- A-4078-BPS. 2. Protein components of the ryanodine receptor complex traffic directly from rough ER to concentrate in junctional SR. Sleiman, N.H, Jones, L., Cala, S. 57th Annual Meeting of the Biophysical Society, Philadelphia, PA, 2013, 13-A-3508-BPS 3. Calsequestrin accumulates around the nucleus in canine heart failure. Sleiman, N., Jacob, S., Sala-Mercado, J., McFarland, T., Sabbah, H., Cala, S. 2nd Annual UM-WSU Joint Physiology Symposium, Ann Arbor, MI Aug 2011. 4. A high-turnover phenotype of calsequestrin with elevated rates of synthesis and degradation replaces normal calsequestrin in heart failure, Sleiman, N., Jacob, S., Sala-Mercado, J., McFarland, T., Sabbah H., Cala, S. 55th Annual Meeting of the Biophysical Society, Baltimore, MD, 2011, 11-A-1772-BPS. 5. The cytosolic protein kinase CK2 is the physiological cardiac calsequestrin kinase, McFarland, T., Sleiman, N., Cala, S. 55th Annual Meeting of Biophysical Society, Baltimore, MD, 2011,11-A-1727-BPS. 6. Cardiac hypertrophy causes newly-synthesized calsequestrin to remain around the nucleus in rough endoplasmic reticulum, Cala, S., Jacob, S., Sleiman, N., Sabbah, H., 54th Annual Meeting of the Biophysical Society, San Francisco, CA, Mar 2010, #10-A-945-BPS.