Characterization of TRP Ion Channels in Cardiac Muscle a Dissertation

Characterization of TRP Ion Channels in Cardiac Muscle

A dissertation submitted

to Kent State University in partial

fulfillment of the requirements for the

degree of Doctor of Philosophy

Spencer R. Andrei

May 2017

Except for previously published materials Dissertation written by

Spencer R. Andrei

B.S., University of Mount Union, 2012

Ph.D., Kent State University, 2017

Approved by

______, Chair, Doctoral Dissertation Committee Derek S. Damron, Ph.D.

______, Member, Doctoral Dissertation Committee Ian N. Bratz, Ph.D.

______, Member, Doctoral Dissertation Committee Colleen Novak, Ph.D.

______, Member, Doctoral Dissertation Committee Soumitra Basu, Ph.D., MBA

______, Graduate Faculty Representative Hanbin Mao, Ph.D.

Accepted by

______, Director, School of Biomedical Sciences Ernest J. Freeman, Ph.D.

______, Dean, College of Arts and Sciences James L. Blank, Ph.D. Table of Contents

LIST OF FIGURES……………………………………………………………………...v

LIST OF TABLES……………………………………………………………………..vii

LIST OF ABBREVIATIONS…………………………………………………………viii

ACKNOWLEDGMENTS………………………………………………………..…..….x

CHAPTER ONE: BACKGROUND...... ……………………………………………...1

Heart Failure Epidemiology…………………………………………………….1

Contractile Machinery of the Heart……………………………………………2

The Cardiac Cycle………………………………………………………3

Ventricular Cardiomyocytes……………………………………………6

Cross-Bridge Cycling and the Sliding Filament Theory……………7

Excitation-Contraction Coupling…………………………………….10

2+ [Ca ]i and Myofilament Sensitivity in Myocardial Contractility

Regulation…………………………………………………………..…16

Heart Failure Pathophysiology…………………………………………..….17

Current Treatment Modalities of Heart Failure…………………….21

TRP Ion Channels Super Family……………………………………………22

TRPA1………………………………………………………………….25

TRPV1……………………………………………………………...…..26

TRPA1 and TRPV1 Interactions………………………………...…..26

TRP Channels and the Cardiovascular System………...………..27

iii

Summary of TRPA1 and TRPV1 in Heart Failure…………………29

CHAPTER TWO: TRPA1 is functionally co-expressed with TRPV1 in cardiac muscle: Co-localization at z-discs, costameres and intercalated discs...…31

Introduction..……………………………………………………………………31

Materials and Methods………………………………………………………..34

Results………………………………………………………………………….41

Discussion………………………………………………………………………56

CHAPTER THREE: Stimulation of TRPA1 and TRPV1 Ion Channels Increase Intracellular Ca2+ Transients and Contraction in Mouse Ventricular Myocytes……………………………………………………………………………….65

Introduction.…………………………………………………………………….65

Materials and Methods ……………………………………………………….67

Results………………………………………………………………………….72

Discussion…………………………………………………………………...…96

CHAPTER FOUR: The role of TRPA1 in myocardial infarction (MI) and ischemia-induced cell death……………………………………………………...103

Introduction…………………………………………………………………...103

Materials and Methods………………………………………………………105

Results………………………………………………………………………...110

Discussion…………………………………………………………………….118

CHAPTER FIVE: CONCLUSIONS………………………………………………...127

REFERENCES……………………………………………………………………….129

List of Figures

Figure 1. Wigger’s diagram…………………………...…………………………...…5

Figure 2. Structural Arrangement of Contractile Filaments in a Cardiac Myofibril and Sarcomere……………………………………………………………..8

Figure 3. Myosin Cross-bridge Cycling During a Normal Contraction Cycle…..11

Figure 4. Ca2+ Cycling During Contraction and Relaxation in a Cardiomyocyte…………………………………………………………….14

Figure 5. A Topological Structure of TRP Channels……………………………..24

Figure 6. TRPA1 and TRPV1 are expressed in CMs obtained from wild-type (WT) mice………………………………………………………………….42

Figure 7. TRPA1 and TRPV1 colocalize throughout the different layers of cardiac muscle………………………………………………………….…………..44

Figure 8. TRPA1 and TRPV1 localize at the costameres and Z-discs in cardiac myofibers…………………………………………………………….…….45

Figure 9. TRPA1 and TRPV1 colocalize at the Z-disc, costameres and intercalated discs in CMs………………………………………….……..48

Figure 10. TRPA1 and TRPV1 stimulation elicits transient increases in intracellular free calcium concentration in quiescent CMs……….....51

Figure 11. AITC and capsaicin induce dose-dependent increases in intracellular free calcium concentration in WT CMs through mechanisms dependent upon TRPA1 and TRPV1, respectively………………..….54

2+ Figure 12. Allyl isothiocyanate (AITC) increases [Ca ]I and shortening in CMs………………………………………………………………………...73

Figure 13. AITC increases fractional shortening, maximum velocity of shortening and maximum velocity of relengthening in CMs………………………75

2+ 2+ Figure 14. AITC increases peak [Ca ]I and accelerates time to peak [Ca ]I and 2+ the rate of [Ca ]I decay in CMs…….…………………………………...78

2+ Figure 15. Capsaicin increases [Ca ]I and contractile function in CMs…….…...82

2+ Figure 16. AITC has no effect on [Ca ]I and shortening in CMs obtained from TRPA1 null mice……………………………………….………………….85

2+ Figure 17. Capsaicin has no effect on [Ca ]I and shortening in CMs obtained from TRPV1 null mice……………………….……………………………88

2+ Figure 18. Treatment with HC030031 or SB366791 Does Not Alter [Ca ]i Dynamics or Contractile Function in CMs……………………………..91

Figure 19. TRPA1 activation with AITC dose-dependently increases ejection fraction in wild-type murine hearts……………………………………...95

Figure 20. TRPA1 gene deletion leads to exaggerated scar formation following myocardial infarction in mice……………….…………………………..111

Figure 21. TRPA1-/- mice exhibit deteriorated cardiac function following MI………………………………………………………….………………114

Figure 22. AITC attenuates ischemia-induced CM cell death…...... 116

List of Tables

Table 1. Comparison of AITC-, capsaicin- and ISO-induced changes in CM 2+ [Ca ]i and contractile function…………………………………………..93

Table 2. TRPA1-/- mice exhibit deteriorated cardiac function following MI……..114

vii

List of Abbreviations

ACEi – Angiotensin converting enzyme inhibitor

ADP – Adenosine diphosphate

AITC – Allyl isothiocyanate

AngII – Angiotensin II

ATP – Adenosine triphosphate

β-AR – Beta-adrenergic receptor

Ca2+ - Calcium ion

2+ [Ca ]I – Intracellular free calcium concentration

CA – Cinnamaldehyde cAMP – cyclic adenosine monophosphate

CICR – Calcium-induced calcium release

CM – Adult mouse ventricular cardiomyocyte

DRG – Dorsal root ganglion

ECC – Excitation-contraction coupling

ECG – Electrocardiogram eNOS – endothelial nitric oxide synthase

HF – Heart failure

HFpEF – Heart failure with preserved ejection fraction

HFrEF – Heart failure with reduced ejection fraction

ISO - Isoproterenol

K+ - Potassium ion

viii

LAD – left anterior descending artery

LTCC – L-type calcium channel

LV – Left ventricle

MCU – Mitochondrial calcium uniporter

MI – Myocardial infarction

MLC2 – Myosin light chain 2

Na2+ - Sodium ion

NCX – Sodium/calcium exchanger

NO – Nitric oxide

PKA – Protein kinase A

PKCε – Protein kinase C epsilon

PLB – Phospholamban

RAAS – Renin-angiotensin-aldosterone system

RYR – Ryanodine receptor

SERCA – Sarcoplasmic reticulum calcium ATPase

SNS – Sympathetic Nervous System

SR – Sarcoplasmic reticulum

Tn(C, I, T) – Troponin (C, I, T)

TRPA1 – Transient receptor potential ankyrin channel subtype-1

TRPA1-/- - TRPA1 knockout

TRPV1 – Transient receptor potential vanilloid channel subtype-1

TRPV1-/- - TRPV1 knockout

WT – wild-type

Acknowledgments

This dissertation would not have been possible without the tremendous support I have received over the course of the past several years. First, I would like to thank my advisor, Dr. Derek Damron. I am truly appreciative and incredibly fortunate to have served as an understudy of Dr. Damron. I am grateful for his valuable insight, guidance, criticisms and support throughout my doctoral studies.

I would also like to thank Dr. Ian Bratz. Dr. Bratz has served as an extraordinary source of knowledge, advice and guidance over the course of the past few years.

I will never be able to put into words the amount of respect I have for both of these men, but this short paragraph will have to suffice. They have prepared me extensively for my future endeavors and have placed me on a trajectory where failure is not an option. I will forever be thankful for the mentorship and friendship of both Dr. Damron and Dr. Bratz.

I’d also like to send my sincerest thanks and appreciation to the members of my doctoral committee, Dr. Colleen Novak and Dr. Soumitra Basu, for their time and energy, as well as their valuable insight into our research and willingness to collaborate. I’d like to thank past and present members of the

Damron and Bratz labs including Dr. Pritam Sinharoy, Dr. Daniel Dellostritto, Dr.

Loral Showalter, Monica Ghosh and John Kmetz for the amazing experience I’ve had in my doctoral studies. I’d also like to thank Dr. Gary Meszaros, Dr. Charles

Thodeti, Dr. Daniel Luther, Dr. Roslin Thoppil, Dr. Holly Cappelli and Ravi

Adapala for valuable experience in microsurgery and associated procedures. I would like to acknowledge the Faculty of Biological and Biomedical Sciences and

Kent State University for funding my doctoral work without which none of this could be possible.

Last, and certainly not least, I’d like to thank my friends and family for their unconditional love, support and sacrifice. I am truly grateful for my mom, dad, sister and nephew who are my biggest supporters and will always be my inspiration to do great things. To my friends, I’d like to express my appreciation for their patience, faith and for dealing with my moodiness when research stressed me out. This dissertation is a dedication to my family, friends, mentors, colleagues and everybody who has helped me become the person I am today.

Thank you.

CHAPTER ONE

BACKGROUND

Heart Failure Epidemiology

The American College of Cardiology defines heart failure (HF) as a

“complex clinical syndrome that results from any structural or functional cardiac disorders that impair the ability of the ventricles to fill with or eject blood” (Hunt et

al., 2009). HF is a syndrome of epidemic proportions associated with high

morbidity and mortality rates, as well as increasingly prevalent re-hospitalization

rates. More than 1-2% of the world population is burdened by HF and this

number is expected to increase exponentially within the next ten years given the

aging of populations. Treatment for this patient population costs our nation an

estimated $30 billion annually; therefore, the cost of HF is high and remains a

significant concern for the healthcare community as it relates to the cost of

national healthcare expenditures.

HF encompasses a wide range of cardiac pathologies, but it can be

generally characterized as ventricular dysfunction that leads to inadequate blood

flow circulation. Furthermore, HF is accompanied by sympathoexcitatory

1 reflexes initiated via its onset and their subsequent injurious tendencies. These reflexes are initially compensatory; however, prolonged stimulation of the sympathetic nervous system (SNS) has been shown to exacerbate the deleterious effects of cardiac injury as it pertains to diminished cardiac function.

In fact, current treatment modalities of HF are aimed at inhibiting neurohormonal

(SNS) hyperactivation; hence, these treatment regimens are designed to decrease cardiac workload (and therefore limiting the energy requirement of the diseased myocardium), decreasing afterload and controlling blood pressure by limiting fluid retention. Although there have been some recent advances in treatment regimens, nearly half of HF patients will die within five years of diagnosis. Therefore, identification of novel therapeutic strategies to combat the development and progression of HF, as well as the underlying diseased contractile machinery, are of the utmost importance and great clinical significance.

Contractile Machinery of the Heart

The heart is a phenomenal organ that consistently pumps throughout a person’s lifetime. It acts as a syncytium by which it rhythmically contracts and relaxes to propel blood through the systemic vasculature to deliver oxygen and nutrients to the organs of the body. Although the heart consists of several different cell types, this dissertation will focus on the contractile cells of the myocardium known as cardiomyocytes (CMs). Millions of CMs are interconnected through a complex network to allow for the heart to contract in a

2 coordinated fashion. In order to understand the role of CMs cardiac physiological process, one must first understand the cardiac cycle.

The Cardiac Cycle

The events occurring in the heart from the beginning of one heartbeat to the beginning of the next is known as the cardiac cycle. Each heartbeat is initiated by an action potential generated in the sinoatrial node located in the right atrium. The action potential travels through the atria and then through the A-V bundle to the ventricles. A slight delay in the passage of the action potential from the atria to the ventricles allows the atria to contract first, effectively emptying blood into the ventricles prior to ventricular contraction.

The cardiac cycle is defined in terms of chamber relaxation and contraction, or diastole and systole, respectively. Furthermore, the cycle can be summarized in an 8-step process; initially, all four chambers are relaxed and are partially filled with blood in the diastasis phase. Next, atrial systole occurs and completes ventricular filling. As atrial systole ends and atrial diastole begins, ventricular contraction occurs and closes the left AV valve in a process known as isovolumetric contraction. ‘Isovolumetric’ refers to the contraction of a chamber without actually propelling blood forward. This process occurs in the ventricles when the amount of pressure generated by ventricular tissue closes the AV valves, but is not great enough to push through the semilunar valves. The second phase of ventricular systole occurs when ventricular pressure exceeds

3 that of the arteries, the semilunar valves open and blood is forced out of the ventricle in a process known as ventricular ejection. Immediately following the cessation of ventricular systole, the semilunar valves will close and ventricular diastole will begin. In early ventricular diastole, blood flows into the relaxed atria but the AV valves remain closed in a process known as isovolumetric relaxation.

In late ventricular diastole, all chambers of the heart are relaxed and begin to fill passively with blood prior to the initiation of the next cardiac cycle. Altogether, a cardiac cycle is known to begin at the initiation of atrial systole and cessation of one cycle is complete at the end of ventricular filling. A summary of the events occurring in the cardiac cycle are shown in Figure 1.

When discussing the phases of the cardiac cycle, one must understand the corresponding electrocardiogram (ECG) indices. Generally, ECGs are defined in terms of ‘waves’ which are electrical voltages generated by the heart.

The ‘P’ wave is caused by atrial depolarization and is followed by atrial systole shortly thereafter. The ‘QRS’ complex occurs during ventricular depolarization immediately prior to the onset of ventricular systole. ‘T’ waves represent ventricular repolarization and subsequent relaxation. In general, the electrical voltages observed in ECGs are indicative of atrial and ventricular depolarization and repolarization patterns. Indeed, any irregularities in cardiac rhythms

(arrhythmias, fibrosis, etc.) can be assessed via ECG analysis. Irregularities in cardiac rhythms often stem from underlying issues within CMs

Figure 1: Wigger’s diagram. The schematic diagram above illustrates the

electro-mechanical relationship that governs the beating of the heart. Taken and modified from Pearson Education, Inc.

5 and their functional properties. In the next section, we will examine the structure, function and beat-to-beat physiology of ventricular CMs in detail (Guyton and

Hall, 2007).

Ventricular Cardiomyocytes

CMs are rod-shaped, cylindrical cardiac muscle cells that range in length from ~100-150 µm long and are specially designed to receive and propagate action potentials forward. Adjacent CMs are connected by structures known as intercalated discs which serve to structurally anchor cells together and to electrically connect them to each other forming a functional syncytium. The intercalated discs are absolutely necessary in order to allow the heart to rapidly beat in a coordinated fashion. The precise mechanisms by which CMs are connected allow the heart to contract in a manner resembling a “wringing” motion whereby the apex of the heart ejects blood toward the base and out of the aorta.

In this dissertation, we isolate ventricular CMs from adult mice and perform a myriad of experiments including measurements of intracellular free calcium

2+ concentration ([Ca ]i) and contractile function during in vitro electrical pacing. In order to understand the process by which CMs contract and relax, one must first understand the basic structure of cardiac muscle.

The functional units of cardiac muscle cells are referred to as sarcomeres.

Upon contraction, these sarcomeres shorten through an intricate mechanism based upon theoretical principles of sliding filaments. Sarcomeres consist of

dark (A) bands and light (I) bands that exhibit a striated appearance when

observed under a microscope. The region that connects the end of one sarcomere to the beginning of the next is referred to as the Z-disc. Repeated sections of sarcomeres compose the tubular CMs and myofibrils. The basic structure of a sarcomere is depicted in Figure 2. The following sections will describe the precise mechanisms underlying muscular contraction and subsequent relaxation.

Cross-Bridge Cycling and the Sliding Filament Theory

Each CM is comprised of complex contractile machinery that serves to carry out the contraction process itself. Within each CM are contractile fibers

known as myofibrils. These myofibrils are composed of two muscular filaments

known as actin and myosin that are responsible for the machinery underlying the

contraction process.

One myosin filament is composed of four light chains and two heavy

chains which are connected to the Z-discs through titin. Two heavy chains

wrapped spirally around each other make up the ‘tail’ of the myosin molecule,

whereas a globular polypeptide structure makes up the myosin ‘head’. The

‘heads’ of the myosin molecule hang outwards to the sides of the filament and

are attached to the double helix by portions referred to as the ‘arm’. Together,

the protrusions made up of the ‘arm’ and ‘head’ are called the cross-bridge.

Figure 2: Structural Arrangement of Contractile Filaments in a Cardiac Myofibril

and Sarcomere. Sarcomeres, the functional units of myofibrils, are composed of

an overlapping arrangement of thick and thin filaments that provide the structural

framework for contraction to occur. This arrangement divides sarcomeres into zones (H-zone and zone of overlap) and distinctive bands (A band and I band).

The z-discs define the end of each sarcomere. Z-discs are located on the end of each thin filament. Structural filaments called titin provide a structural connection between the z-discs and thick filaments. The M line is an arrangement of proteins that defines the middle of the sarcomere.

A notable feature of the myosin head is that it functions as an ATPase enzyme,

which is critical for muscle contraction – a phenomenon that will be discussed in

the upcoming paragraphs.

The actin filaments are composed of three protein components: actin,

tropomyosin and the troponin complex. Similarly to the myosin molecules, the backbone of actin is composed of two helical strands of F-actin and tropomyosin molecules. In the resting condition, tropomyosin molecules cover the active sites of actin to block the interaction between actin and the myosin head. Each tropomyosin molecule has an attached troponin complex that contains three loosely bound protein subunits: troponin I (TnI), troponin T (TnT), and troponin C

(TnC). TnI, TnT and TnC have particularly strong affinities for actin, tropomyosin and calcium ions, respectively. However, the affinity of TnC for calcium is the primary determinant for the initiation of the contraction process itself (Bers,

1997).

The interaction between actin and myosin during muscular contraction is initiated by the action potential-induced release of calcium from a mesh-like network surrounding the cardiac myofibrils known as the sarcoplasmic reticulum

(SR). Under normal resting conditions, the troponin-tropomyosin complex inhibits the active sites on the actin filaments. Upon its release from the SR, Ca2+

binds TnC which induces a conformational change that essentially tugs the tropomyosin molecule away from the active site of actin, thus relieving the inhibitory effect of the troponin-tropomyosin complex. At this stage, the actin

filament can be considered ‘activated’. Upon actin filament activation, the heads

of the myosin cross-bridges bind actin in multiple sites based upon the relative

Ca2+ availability in the myofibrillar network. Upon binding to actin, the myosin heads tilt toward the arm and drags the actin filaments alongside in a

phenomenon referred to as the power stroke (Figure 3). The myosin heads then

detach and either rebind the actin to induce another power stroke or remain

unattached to allow for relaxation. The process of the power stroke, however,

requires ATP. ATP binds the myosin heads prior to contraction and becomes

hydrolyzed to ADP plus Pi. Once the myosin head enzymatically hydrolyzes

ATP, it stores the energy which is used to induce the sliding-filament mechanism.

After contraction, the ADP and Pi are released for a new ATP to bind which

allows another contraction to ensue (Bers, 1997). The process described above

is completely dependent upon the presence of Ca2+ and its essential role in

muscular contraction through a process known as excitation-contraction coupling

(ECC).

Excitation-Contraction Coupling

ECC refers to the series of events beginning with an electrical action potential

and its subsequent initiation of muscle cell contraction (Bers, 1997) – a process

in which calcium handling is absolutely critical (Marks, 2013). In CMs, electrical

excitation travels along the sarcolemmal membrane which propagates as a wave

of depolarization along the surface and the transverse tubules (T-tubules). The

mechanism by which the action potential propagates allows for

Figure 3: Myosin Cross-bridge Cycling During a Normal Contraction Cycle.

Myosin cross-bridge formation is under the regulation of ATP hydrolysis by the

ATPase on the myosin head. Initially, the myosin heads hydrolyze ATP and

become “charged”. Second, the myosin binds to actin to form the cross-bridge.

Next, the myosin head releases ADP which results in a conformational change in

the protein that causes contraction (power stroke). Lastly, myosin heads bind the bioavailable ATP and detach from actin and the contraction cycle will continue as long as ATP and Ca2+ is available.

rapid transmission of electrical impulses to each muscle fiber. The wave of

depolarization subsequently causes opening of the voltage-gated calcium

channels, commonly referred to as L-type calcium channels (LTCC), which allow

Ca2+ influx down its concentration gradient and into the intracellular cytosolic space. Notably, the amount of Ca2+ that enters through the LTCC is not, in itself,

enough to trigger muscle contraction. However, the LTCC-mediated Ca2+ influx

stimulates Ca2+ release from the SR through an intricate mechanism necessary to induce muscular contraction (Bers, 2001).

The SR stores massive amounts of Ca2+ and is a major regulator of

myocardial contraction. In fact, part of the SR is located adjacent to the T-

tubules for efficient calcium-mediated activation of the ryanodine receptor (RYR) on the SR; this stimulation of RYR allows for large amounts of Ca2+ to be

released from the SR in a process known as calcium-induced calcium release

(CICR). The Ca2+ released from the SR enters the cytosolic space and is used to

initiate actin and myosin cross-bridge cycling by binding to TnC on the actin thin

filaments, moving tropomyosin aside to allow for the sliding filament mechanism

and contraction to ensue (Bers, 2001).

Additionally, the mechanisms by which cardiac muscle undergoes diastole

are of equal importance. In order for cardiac muscle to relax, the intracellular

2+ 2+ free calcium concentration ([Ca ]i) must be decreased so Ca releases TnC and

allow the myofibrils to relax. This removal of Ca2+ from the cytosolic space

following a muscle contraction is typically accomplished through an intricate

system of ion pumps and transporters including SR Ca2+-ATPases (SERCA),

sarcolemmal Na2+-Ca2+ exchangers (NCX), sarcolemmal Ca2+-ATPases and the mitochondrial Ca2+-uniporters (MCU). The majority of studies conducted in

mammalian ventricular tissue demonstrate that SERCA is primarily responsible

for the majority of Ca2+ removal from the cytosolic space, with the NCX being

responsible for a little less than 10% (Bers, 2001). SERCA is located on the SR

membrane and serves to pump Ca2+ back into the sarcoplasmic reticulum in

between each contraction – effectively “loading” the SR for the next action potential-induced stimulus. While explaining the function of SERCA, one must

include the closely associated inhibitory protein called phospholamban (PLB). In

the basal state, PLB inhibits the action of SERCA pumps; however,

phosphorylation of PLB removes the inhibition and allows SERCA to pump Ca2+

back into the SR (Bers, 2001). Aside from the SERCA pump, NCX is also

involved in Ca2+ removal and is classified as a transporter located on the

sarcolemma that transports one Ca2+ ion from the cytosolic space to the

extracellular space while simultaneously transporting three sodium (Na2+) ions

into the cell. The primary, or “forward”, mode of this transporter functions as

described above; however, when concentration gradients vary from homeostatic

conditions, this transporter is able to become reversible and operates in “reverse”

2+ mode by which it will increase [Ca ]I (Bers, 2001). Indeed, proper functioning of

these transporters, pumps and SR-mediated Ca2+-handling is essential for the

contractile machinery of cardiac muscle to operate efficiently. This process is

demonstrated in Figure 4.

Figure 4: Ca2+ Cycling During Contraction and Relaxation in a Cardiomyocyte.

Propagation of the action potential into T-tubules activates voltage-gated Ca2+

channels (L-type Ca2+ channels) causing influx of Ca2+.from the extracellular

space into the intracellular compartment. The influx of Ca2+ activates ryanodine

receptors (RYR) located on the sarcoplasmic reticulum (SR) which initiates the

release of Ca2+ into the cytosol. This process of Ca2+ influx triggers the release

of stored Ca2+ and is referred to as Ca2+-induced Ca2+-release (CICR). This is a characteristic feature of cardiac muscle. The Ca2+ release from the SR then initiates molecular mechanisms of contraction by binding to TnC, as mentioned earlier. In order for relaxation to ensue, Ca2+ needs to be removed from the

cytosol. The SR Ca2+ ATPase (SERCA) actively pumps Ca2+ back into the SR.

The Na+-Ca2+ exchanger (NCX) also participates in removing Ca2+ from the

cytosol into the extracellular space. The Ca2+ uniporter in the mitochondrial

membrane (MCU) and a Ca2+-ATPase on the sarcolemmal membrane also function to remove Ca2+ however they have limited roles.

2+ [Ca ]i and Myofilament Sensitivity in Myocardial Contractility Regulation

There are two predominant mechanisms that regulate CM contractility: 1)

2+ [Ca ]i handling, as described above and 2) the sensitivity of myofilaments to

Ca2+. The latter is part of a mechanism in which the CMs can alter the intrinsic force of contraction in response to the amount of incoming blood. Also, CM contractility is intrinsically manipulated in order to meet the changes in metabolic demand of the myocardial tissue. As previously described, Ca2+ influx from the

extracellular space and SR are responsible for driving contractile machinery. The

same pumps and transporters described earlier play roles in Ca2+- clearance during diastole and are under direct regulation of intracellular protein kinases that

2+ modulate [Ca ]i and contractility.

Myofilament Ca2+ sensitivity has recently garnered significant interest

among clinicians, researchers and pharmaceutical companies as it pertains to

the regulation of myocardial contractility. The sensitivity of myofilaments to Ca2+

is generally considered to be primarily managed by phosphorylation of TnI, which

is located on the thin filament, although myosin light chain 2 (MLC2) and TnT

have also been implicated in the process, as well (Monasky et al., 2013).

Furthermore, intracellular pH has been demonstrated to be a critical regulator of

alterations in myofilament Ca2+ sensitivity where increases in intracellular pH

levels (alkalosis) increases myofilament Ca2+ sensitivity and decreases in pH

levels (acidosis) decrease the sensitivity of myofilaments to Ca2+. Moreover, the

proteins involved in altering myofilament Ca2+ sensitivity are well understood to be downstream mediators of protein kinase A (PKA). As such, PKA-mediated

TnI phosphorylation results in a decreased affinity of TnC for Ca2+ and the

consequential increased rate of dissociation results in a positive lusotropic

phenomenon (Bers, 2001).

2+ Indeed, maintaining homeostatic levels of [Ca ]i and myofilament sensitivity is crucial to the normal functioning of the myocardium. However, pathological conditions, such as those observed in HF, modulate the

2+ mechanisms that regulate CM [Ca ]i handling and contractile function.

Heart Failure Pathophysiology

HF can develop from any number of comorbidities in a patient which

include myocardial infarction, coronary artery disease, hypertension, left

ventricular remodeling and hypertrophied ventricular tissue, among many others

(Metra et al., 2017). Historically, HF has been associated with a reduced ejection

fraction (HFrEF); however, recent epidemiological evidence suggests that the

age adjusted mortality for cardiovascular diseases and coronary artery disease is

decreasing in populations around the globe (Pearson-Stuttard et al., 2016;

Townsend et al., 2015). This is most likely due to better prevention regimens as

well as percutaneous interventions for acute coronary syndromes (De Luca et al.,

2015). As such, a change in clinical characteristics has been observed in

patients who develop HF. The main phenotype of patients with HF is now related

to prolonged hypertension and hypertrophied, poorly relaxing ventricular tissue

leading to a condition known as HF with preserved ejection fraction (HFpEF).

Therefore, HFrEF and HFpEF are used interchangeably with systolic HF and diastolic HF, respectively. Although characteristics historically associated with traditional types of HF are commonly observed in patients with HFpEF

(myocardial fibrosis and stiffness, calcium instability, modulation of contractile proteins, etc.), the precise underlying mechanisms have not yet been elucidated and no specific therapy has been employed. Although it remains important for future investigations to target the pathological conditions underlying HFpEF independently, this dissertation will have a broader focus on HF (including both

HFrEF and HFpEF).

As cardiac output falls and the resultant inability to provide systemic blood flow ensues, a myriad of systemic effects take place in an attempt to restore the heart to its normal working condition to deliver nutrient rich blood in order meet the metabolic demands of the body. The most well-recognized of the compensatory mechanisms stimulated to combat the reduced pumping capacity of the heart is neurohormonal activation which includes activation of the SNS and the renin-angiotensin-aldosterone system (RAAS). Neurohormonal activation constitutes responses that are initiated to maintain cardiovascular homeostasis.

SNS and RAAS activation is initiated to restore cardiac output through processes that: 1) increase the retention of water and salt, 2) increase contractility, 3) induce peripheral arterial vasoconstriction and 4) modulate the release of inflammatory mediators that promote repair and remodeling of cardiac tissue

(Hartupee and Mann, 2017).

In the short term, these neurohormonal activation mechanisms are

compensatory and beneficial to restore homeostasis in the heart, kidneys, and

vasculature; however, long term stimulation of these neurohormonal mechanisms

drives deleterious effects on the cardiovascular system that further exacerbate

the progression of HF by inducing left ventricular (LV) stiffness, increased afterload and poorly relaxing ventricles.

Although the cellular signal transduction pathways underlying HF are subject to extensive investigations, the precise mechanistic details, as well as their pathophysiological implications, remain to be fully determined. However, it is well known that the sustained elevation of SNS activity in HF initializes cellular and molecular modifications that eventually lead to the development of left ventricular remodeling (Hartupee and Mann, 2017). These alterations observed in LV remodeling may include changes in CM biology and energetics that lead to

CM hypertrophy, β-adrenergic receptor (β-AR) desensitization, transcriptional reprogramming of CMs, modified ECC mechanisms as well as altered composition of intracellular and extracellular components (Hartupee and Mann,

2017). In the following paragraphs of this dissertation, we will analyze the modifications of CM ECC and β-AR responsiveness.

As previously stated, ECC refers to the series of events beginning with an electrical action potential and its subsequent initiation of muscle cell contraction.

In the failing heart, the processes by which calcium is mobilized through transporters and pumps are drastically altered. In fact, the manipulations in calcium handling observed in the failing heart are major determinants for the

consequentially diminished systolic and diastolic function. The principal

phenomenon underlying impaired Ca2+ transit in CMs begins with the depleted

Ca2+ levels available in the SR. Hyperphosphorylated RYR results in

destabilization of the closed-state RYR which leads to diastolic Ca2+ leak (Marx

et al., 2000). Moreover, SERCA expression and function are decreased in the

failing heart which further exacerbates the depleted SR Ca2+ bioavailability by inhibiting Ca2+ reuptake (Arai et al., 1993; Hasenfuss et al., 1994). This SERCA

dysfunction is caused in part by a down-regulation of phospholamban

phosphorylation which induces an increased inhibitory effect of phospholamban

on SERCA pump activity. Although a myriad of pathophysiological indices are

observed in failing CM ECC dynamics, several novel treatment regimens appear

to be targeting SERCA and RYR to reestablish their proper homeostatic

functioning (Francis et al., 2014). We will discuss these treatment modalities in

the upcoming section of this dissertation.

Parallel events develop in the failing heart that affects β-AR

responsiveness most of which can be attributed to sustained SNS activity

(Lymperopoulos et al., 2013). Most of the changes in β-AR responsiveness are

generally attributed to three major mechanisms: 1) down-regulation of β1- receptors, 2) β1-receptor dysfunction and 3) β1-receptor desensitization, which

ultimately uncouples the receptors from their downstream G-protein signaling cascade (Port and Bristow, 2001; Rockman et al., 2002). In fact, a decrease in

β-receptor-mediated adenylyl cyclase stimulation and contractile responses were initially detected in the failing myocardium in 1982 (Bristow et al., 1982). The

20 treatment regimens designed to alleviate these affects will be discussed in the upcoming section.

Current Treatment Modalities of Heart Failure

Current therapeutic strategies for treating patients with HF have remained unchanged for nearly 30 years. Although these treatment modalities exhibit some clinical benefit in reducing morbidity and mortality, most patients will succumb to the devastating disease within 5 years of diagnosis. The strategy today is to prescribe β-adrenergic receptor blockers in combination with angiotensin converting enzyme (ACE) inhibitors or angiotensin II (AngII) receptor antagonists, as well as diuretics or aldosterone receptor antagonists. The strategy behind this combination therapy is to minimize the deleterious effects observed with SNS hyperactivity. β-blockers are used to alleviate unwanted effects on heart rate and blood pressure; this will reduce the “load” and energy expenditure which ultimately reduces the increased pressures observed at elevated heart rates. ACE inhibitors and AngII antagonists are used to mitigate the hypertensive effects of SNS activity whereas diuretics and aldosterone receptor antagonists lower salt and water retention in an effort to reduce blood volume and pressure.

Theoretically, positive inotropic therapy would increase the diminished cardiac function observed in the failing myocardium; however, the vast majority of clinical studies have demonstrated an actual worsening of prognosis and re- hospitalization rates in patients placed on inotropic support (Francis et al., 2014).

Therefore, positive inotropes have only been indicated in severe septic and

cardiogenic shock (acute phase), severe systolic failure, bridge to heart

replacement therapy and palliative care (Francis et al., 2014). The current

strategy that is employed to increase cardiac inotropy in the setting of

combination therapy is through the use of cardiac glycosides such as digitalis.

Digitalis compounds are Na+/K+-ATPase inhibitors which lead to elevations of intracellular Na+ levels. The increased Na+ concentration then triggers reverse

mode NCX activity, effectively extruding Na+ out of the cell in exchange for Ca2+

2+ to come into the CM. The increase in [Ca ]i induces CICR, thereby making more

Ca2+ available to bind to troponin-C and increase contractility. Although this

increase in contractility would theoretically manifest itself as significant changes

in overall cardiac output, ejection fraction is typically increased by less than 10%

(Francis et al., 2014). Furthermore, other concerns with digitalis include

cytotoxicity at optimal concentrations, altered CM repolarization and minimal

effects on re-hospitalization rates (Francis et al., 2014).

Taken together, the current therapeutic strategies aimed at combating HF provide minimal benefits that seem to reduce unwanted side effects instead of reversing the debilitating disorder. As such, there is a dire need for the development of novel treatment modalities and pharmaceutical agents to counteract the deleterious ramifications observed in patients burdened by HF.

TRP Ion Channels Super Family

The cell membrane is a crucial regulator of bidirectional transport whereby it’s permeability to ions (potassium, calcium, sodium, etc.) is dependent upon its concentration gradient and cellular homeostasis. Many physiological

mechanisms depend on the structural integrity of the membrane and its’ inherent

ability to modulate permeability of substances based upon intracellular and

extracellular environments. Embedded within the membrane are several types of

ion channels responsible for the transport of substances. The mechanisms by

which these ion channels modulate cellular events are subject to continuous

investigations and are consistently implicated in several physiological and

pathophysiological events. Furthermore, the cellular signal transduction

cascades elicited via TRP channel stimulation have gained recent attention and

indicate that these ion channels may serve as targets for therapeutic

interventions in a myriad of pathological disorders.

Transient Receptor Potential (TRP) channels have rapidly emerged as attractive targets for clinical intervention. TRP channels are structurally-related, non-selective cation channels characterized by their high permeability to calcium

(Fernandes et al., 2011). The TRP superfamily currently consists of 28 members categorized into 7 subfamilies: TRPC (canonical), TRPV (vanilloid), TRPM

(melastatin), TRPA (ankyrin), TRPP (polycystic), TRPML (mucolipin) and, a gene observed only in non-mammals, TRPN. TRP channels are key detectors of

noxious stimuli and can sense a variety of other stimuli including shear stress,

mechanical stress, inflammation products and pressure, among others

(Clapham, 2003; Dhaka et al., 2006; Julius and Basbaum, 2001). A topological

structure of a TRP channel is shown in Figure 5.

Figure 5: A Topological Structure of TRP Channels. A subunit of TRP channels

has six transmembrane (TM) domains. A pore-forming loop is located between

the TM5 and TM6 domains to allow cation influx. N- and C-termini are located in the cytosolic space. The amino terminus is the location of ankyrin repeats in

some TRP channels. Taken and modified from Hwang 2014.

The primary structures of TRP channels predict six transmembrane

domains (S1-S6) where the channel pore is located between S5 and S6. These

pores open in response to channel activation, most of which are suggested to be

polymodal in nature. This particular feature of TRP channels confers diverse

physiological and pathophysiological involvement in cellular events. In the

upcoming subChapter, two members of the TRP superfamily will be introduced

and described: TRPA1 and TRPV1.

TRPA1

TRPA1 is characterized by its 14-19 ankyrin repeats at its N-terminus and

was first cloned from lung fibroblasts in 1999 by the Jaquemar lab (Bessac and

Jordt, 2008; Jaquemar et al., 1999; Story et al., 2003). Previous investigations

suggest that the numerous ankyrin repeats at the N-terminus are essential for modulating channel activity, receptor sensitivity and protein-protein interactions

(Nilius et al., 2001). TRPA1 is predominantly expressed in non-myelinated C fibres of dorsal root ganglion (DRG) neurons, which are sensors for noxious stimuli and temperature. As such, the channels have been identified in playing crucial roles in nociceptive information transduction. Additionally, TRPA1 expression patterns have been identified in hair cells, nodose ganglia, trigeminal neurons and epithelial cells (Nilius et al., 2012; Stokes et al., 2006) where they can be activated by several exogenous and endogenous compounds including allyl isothiocyanate (mustard oil) (Bandell et al., 2004; Jordt et al., 2004),

25 cinnamaldehyde (cinnamon), allicin (garlic) (Macpherson et al., 2005) and nitric oxide (Miyamoto et al., 2009) among others.

TRPV1

TRPV1 is one of the most investigated ion channels of the TRP channel superfamily. Formerly referred to as VR1 or the ‘capsaicin-receptor’, TRPV1 was first identified as a thermo-sensitive receptor and cloned in 1997 by the Caterina lab (Caterina et al., 1997). Future studies identified the presence of the ion channel in a number of species throughout the central nervous system and small-to-medium sized neurons of DRG (Mezey et al., 2000; Roberts et al.,

2004). Moreover, TRPV1 expression has been demonstrated in smooth muscle cells and endothelial cells (Birder et al., 2001; DelloStritto et al., 2016) where it is able to be activated with high heat (>42ºC), low pH (Caterina et al., 1997), capsaicin (peppers)(McNamara et al., 2005) and allicin (Macpherson et al.,

2005), among several others.

TRPA1 and TRPV1 Interactions

Emerging evidence indicates that TRPA1 and TRPV1 demonstrate reciprocal regulation, suggesting that these channels may cross-talk when both are present in a cell. In fact, recent evidence suggests that nearly 90% of all

TRPA1-expressing neurons also express TRPV1 while only 30% of TRPV1- expressing neurons express TRPA1 (Katsura et al., 2006; Kobayashi et al.,

2005). Previous studies have demonstrated that these channels may interact

indirectly or via direct protein-protein interactions (Akopian, 2011; Sadofsky et al.,

2014; Staruschenko et al., 2010). Cross-talk mechanisms in co-expressed

TRPA1 and TRPV1 have already been demonstrated in human airway epithelial cells, the vascular bed and sensory neurons (Kamakura et al., 2013; Nassini et al., 2012). Previous studies have demonstrated TRPV1-mediated regulation of

TRPA1 sensitivity in response to agonist stimulation (Akopian et al., 2007;

Akopian et al., 2008). In fact, our laboratory has previously demonstrated cross-

talk mechanisms by which TRPA1 and TRPV1 interact in sensory neurons

through an eNOS-mediated, PKCε-dependent signal transduction pathway

(Sinharoy et al., 2015; Wickley et al., 2010; Zhang et al., 2011). However, the

extent to which these channels interact in specific subsets of cell populations and

how they regulate each other’s physiological and/or pathophysiological

functionality remains to be fully elucidated. Furthermore, the precise

ultrastructural localization and function of TRPA1 and TRPV1 ion channels has

not been explored in CMs.

TRP Channels and the Cardiovascular System

The diverse roles of TRP channels in the cardiovascular system were

conveyed in a well-written review by the Yue laboratory (Yue et al., 2015). The

review reports the extensive involvement of the TRP ion channel superfamily in

the cardiovascular system. This dissertation, however, will examine the roles of

only TRPA1 and TRPV1 ion channels in the cardiovascular system.

TRPA1 and TRPV1 expression and function in the cardiovascular system

has predominantly been limited to the vasculature. These ion channels have

been identified in vascular endothelial cells and smooth muscle cells whereby

their activation elicits vasodilation (Bratz et al., 2008; Earley, 2012). Although the

notable potential of these channels in modulating vascular physiology is

promising and warrants further investigations, this dissertation will focus on

examining the roles of TRPA1 and TRPV1 in cardiac muscle, specifically.

Historically, the extent to which TRPV1 mediates cardiac dynamics has

been limited to pathophysiological models, which will be discussed in the

upcoming subChapter. The role of TRPA1 in modulating cardiac function has garnered recent attention, even though the presence of TRPA1 in cardiac tissue has only been demonstrated at the protein level in fibroblasts (Oguri et al., 2014)

and at the mRNA levels in CMs (Pazienza et al., 2014). A recent study suggests

that TRPA1 mediates changes in heart rate variability following exposure to the

TRPA1-agonist, acrolein, in mice (Kurhanewicz et al., 2016). However, a

conflicting investigation conducted by Susan Brain’s laboratory demonstrates that

knockout of TRPA1 did not result in any significant alterations in cardiac function

(Bodkin et al., 2014). Furthermore, the traditional non-selective TRPA1 agonist,

cinnamaldehyde (CA), has been demonstrated to accelerate heart rate, although

this effect was not concluded to work through TRPA1 (Alvarez-Collazo et al.,

2014). In fact, CA was excluded from most the studies in this dissertation due to the implicated role of the compound in inhibiting L-type Ca2+ channels in mouse

ventricular CMs and vascular smooth muscle cells. The relative polymodal

nature of TRPA1 and TRPV1 activation confers involvement in diverse

physiological and pathophysiological mechanisms. As such, identifying the

extent to which the presence and activation of TRPA1 and/or TRPV1 modulates cardiac physiology is of the utmost translational significance.

Summary of TRPA1 and TRPV1 in Heart Failure

The therapeutic potential for TRPA1 and TRPV1 in alleviating the deleterious effects observed in cardiovascular disorders has garnered significant attention across the scientific community over the course of the past decade.

The vast majority of the literature analyzes the extent to which these ion channels regulate vasoactivity, where activation of TRPA1 or TRPV1 has been implicated in treating hypertension (Yue et al., 2015). As such, there is an obvious paucity of information in the literature describing the extent to which

TRPA1 and TRPV1 are involved in mediating physiological cardiac cell processes; this is most likely due to the previously unknown expression patterns of the ion channels throughout cardiac muscle or CMs.

Historically, TRPV1 channels in the heart have been investigated with regards to their roles in regulating pathophysiological events. TRPV1 has repeatedly been implicated in cardioprotection following myocardial infarction and ischemia-reperfusion injury conducted in the heart (Huang et al., 2009), as well as other tissue beds (Chen et al., 2014). Furthermore, a role for TRPV1 in attenuating high salt- and pressure overload-induced cardiac hypertrophy has

been postulated (Lang et al., 2014; Wang et al., 2014). In chronic refractory

angina, inhibiting TRPV1 has been associated with the amelioration of chest

pain; however, stimulation of TRPV1 channel activity has been linked with

vasodilation in hypertension, diminished myocardial injury after heart attacks and

depressed formation of atherosclerotic plaques leading to infarction (Robbins et

al., 2013). Although there is clear evidence for a role of TRPV1 in cardiac

pathophysiology, the extent to which it is involved in basal cardiac physiology has

yet to be fully determined. Apace with TRPV1, a lack of information exists

describing the physiological, as well as pathophysiological, cardiac cell

processes to which TRPA1 channels are involved. In fact, theories involving the

potential role of TRPA1 channels in the heart are highly speculative due to the

previously unknown presence of the protein in CMs.

As stated previously, mishandling of Ca2+ in CMs serves to trigger the

progression of HF, arrhythmias, and cardiac remodeling. As non-selective cation

channels, it has been hypothesized that TRP channels may play significant roles

in mediating these processes (Vennekens, 2011). Particularly, the affinity of

these channels for Ca2+ transport confers a probable role of TRPA1 and TRPV1

in the regulation of myocardial contractility. We tested the hypothesis that

2+ TRPA1 and TRPV1 ion channels will 1) induce transient increases in [Ca ]I upon

2+ stimulation, 2) increase [Ca ]I and cell shortening upon activation in electrically- paced CMs and 3) serve a cardioprotective role in myocardial infarction in mice.

This dissertation will analyze the extent to which TRPA1 and TRPV1 modulate both physiological and pathophysiological cellular events in the heart. The

30 current studies will lay the foundation for the development of novel therapeutic agents designed to not only alleviate the symptoms traditionally associated with

HF, but also to potentially reverse cardiac dysfunction to a healthy homeostatic condition.

CHAPTER TWO

TRPA1 is functionally co-expressed with TRPV1 in cardiac muscle: Co-

localization at z-discs, costameres and intercalated discs

INTRODUCTION

Transient receptor potential (TRP) ion channels of the ankyrin 1 (TRPA1) and vanilloid 1 (TRPV1) subtypes are members of the TRP superfamily of structurally related, non-selective cation channels first described in sensory neurons that are highly permeable to calcium (Fernandes et al., 2011). Recent evidence suggests that TRPA1 and TRPV1 receptors exhibit reciprocal regulation, indicating cross-talk between the two receptors when co-expressed in the same cell (Patil et al., 2010; Staruschenko et al., 2010) and both channels play an important role in the induction of neurogenic pain and inflammation

(Caterina and Julius, 2001; Meseguer et al., 2014; Schwartz et al., 2013).

However, there is accumulating evidence that TRPA1 and TRPV1 have functional roles independent of sensory neurons. In fact, emerging evidence

32 indicates TRPA1 and TRPV1 are expressed in various of other cell types including smooth muscle cells and endothelial cells (Yue et al., 2015).

To our knowledge the expression of TRPA1 at the protein level in cardiac muscle as well as evidence identifying the functionality of the channel and its ultrastructural location in cardiomyocytes (CMs) has yet to be described. One recent study identified the presence of TRPA1 mRNA in mouse CMs (Pazienza et al., 2014), whereas only one other study has demonstrated expression of

TRPA1 at the protein level in cardiac fibroblasts (Oguri et al., 2014). However, several studies conducted over the past decade have demonstrated a cardioprotective role for TRPV1 in the setting of myocardial ischemia and reperfusion injury (Lu et al., 2014) in addition to attenuating high salt-induced cardiac hypertrophy (Lang et al., 2014) and ameliorating pressure overload- induced hypertrophy (Wang et al., 2014). Moreover, TRPV1 channels have recently been described to primarily localize near the epicardial surface of the heart (Zhong and Wang, 2009), however the precise location of the TRPV1 channels within the ultrastructure of the CM has yet to be reported. Since evidence exists in sensory neurons and heterologous expression systems that

TRPA1 and TRPV1 may cross-regulate each other’s function serving as

“partners in crime” (Wickley et al., 2010; Zhang et al., 2011), we questioned whether the potential for a similar paradigm may exist if in fact TRPA1 is coexpressed with TRPV1 in adult mouse CM’s. Uncovering the precise location of TRPA1 and TRPV1 within the ultrastructure of the CM could provide important

information as to the specific role(s) the channels mediate in physiological and/or

pathophysiological events in the heart.

In the current study, we examined the extent to which TRPA1 and TRPV1

are co-expressed in adult mouse CM’s, assessed whether they are expressed

throughout the entire myocardium or localized to specific layers of the heart and

identified their precise ultrastructural location within the isolated CM cytoskeleton.

Moreover, we have explored the extent to which TRPA1 and TRPV1 are

functionally active by assessing changes in intracellular free Ca2+ concentration

2+ ([Ca ]i) in response to increasing concentrations of allyl isothiocyanate (AITC;

specific TRPA1 agonist) or capsaicin (specific TRPV1 agonist) in CM’s obtained

from WT as well as TRPA1-/- and TRPV1-/- mice. Dose response curves for

TRPA1 channel inhibition (HC-030031) or TRPV1 channel inhibition (SB366791)

2+ on the agonist-induced rises in [Ca ]I were also performed in CM’s obtained

from WT mice. The major findings of the current study are that both TRPA1 and

TRPV1 are co-expressed in CMs throughout the endocardium, myocardium and

epicardium and appear to specifically colocalize at the Z-discs and costameres.

2+ Moreover, both TRPA1 and TRPV1 agonists elicit transient rises in [Ca ]i that are absent in CMs obtained from TRPA1-/- and TRPV1-/- mice and attenuated in a

dose-dependent manner in WT CMs pretreated with specific TRPA1 and TRPV1

channel antagonists. The current studies will lay the foundation for future studies

investigating the extent to which cross-talk/regulation between TRPA1 and

TRPV1 play a role in mediating the physiology and pathophysiology of cardiac tissue.

MATERIALS AND METHODS

Animal Model

4-month-old male C57BL/6 mice (n = 6/group) were used and maintained in accordance with the Guide for the Care and Use of Laboratory Animals (NIH).

All animals were housed at the Kent State University animal care facility (Kent,

OH), which is accredited by the American Association for Accreditation of

Laboratory Animal Care.

Isolation of CMs

Murine hearts were excised and transferred to a Langendorff apparatus for CM isolation, as previously described (O'Connell et al., 2007). In brief, mice were sacrificed via cervical dislocation and hearts were rapidly excised then placed into a dish containing perfusion buffer. After the aorta was cannulated and blood was flushed, the hearts were subjected to retrograde perfusion at 37°C and pH 7.4 with a modified Krebs-Henseleit buffer (in mM: 120.4 NaCl, 4.8 KCl, 0.6

KH2PO4, 0.6 Na2HPO4, 1.2 MgSO4-7HsO, 10 Na-HEPES, 4.6 NaHCO3, 30 taurine, 10 BDM, and 5.5 glucose). The calcium-free buffer was sterile-filtered and paced with a peristaltic pump (Masterflex) to begin retrograde perfusion of the heart at a rate of 4 mL/min. After perfusion for 4 minutes, the same solution containing collagenase type II (300 U/mg, Worthington Biochemical) perfused the heart for an additional 8 minutes until the heart became soft. The left ventricles

were removed, minced, then triturated in Krebs-Henseleit buffer containing fetal

bovine serum. The resulting cellular digest was washed and resuspended in

HEPES-buffered saline (in mM: 118 NaCl, 4.8 KCl, 0.6 KH2PO4, 4.6 NaHCO3,

0.6 NaH2PO4, 5.5 glucose, pH 7.4) at 23°C. CM yield was typically ~80-90%.

CMs were then either subjected to immunoblotting, immunocytochemistry, or

2+ slow calcium reintroduction ([1.23 mM]) and subsequent [Ca ]i measurements.

F-11 Cell Transfection with TRPV1 or TRPA1

F-11 cell transfection was carried out as previously described (Sinharoy et

al., 2015). Cultured F-11 cells (hybridoma cell line) were transfected with TRPA1 or TRPV1 cDNA via electroporation using a Neon Transfection System

(Invitrogen). In brief, cultured F-11 cells were harvested and washed with phosphate-buffered saline without calcium or magnesium. The cells were then resuspended in electrolytic buffer, where TRPA1 and TRPV1 cDNA was then added. Using a pulse voltage of 1500 V, a pulse width of 35 msec, and a pulse number of 2, electroporation was then performed. The cells were then suspended in Dulbecco’s Modified Eagle’s Medium supplemented with 10% fetal bovine serum at 37°C. The cells were used to demonstrate the presence of

TRPA1 or TRPV1 in immunoblot analysis.

Preparation of Cell Lysates and Immunoblot Analysis

Immunoblot analysis was performed as previously described (Wickley et al., 2006). CMs were homogenized in a lysis buffer (in mM: 25 Tris-HCl, 150

NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS, pH 7.6) and protein

concentration was assessed using the Bradford method (Bradford, 1976). All

samples were adjusted to 2 mg/mL protein concentration in sample buffer.

Samples containing equal amount of protein lysates (50 µg) were boiled then

subjected to SDS-PAGE on a 4-15% precast polyacrylamide gels (Bio-Rad),

through the use of a minigel apparatus, which were then transferred to

nitrocellulose membranes. Nonspecific binding was blocked with 5% nonfat milk

in Tris-buffered saline solution (0.1% [vol/vol] Tween-20 in 20 mM Tris base, 137

mM NaCl, pH 7.6 containing 3% bovine serum albumin) for 45 minutes at room

temperature. Antibodies against TRPA1 (Novus Biologicals) and TRPV1 (Santa

Cruz Biotechnology) were diluted 1:500 in Tris-buffered saline containing 5% nonfat milk and incubated at 4°C overnight. After washing in Tris-buffered saline, membranes were incubated for 1 h at room temperature with horseradish- peroxidase linked secondary antibody (goat anti-mouse and goat anti-rabbit) diluted 1:5000 in Tris-buffered saline with 5% nonfat milk. Antibody detection was conducted via enhanced chemiluminescence with an ImageQuant LAS 4000 Mini

(General Electric) and immunoreactivity was quantified by scanning densitometry and analyzed using ImageJ software (NIH).

Immunocytochemistry

Immunocytochemistry techniques were carried out as previously described (Wickley et al., 2006). CMs were allowed to adhere to laminin-coated

coverslips (20 µg/mL) at 37°C for 2 h and subsequently fixed in a 1:1

acetone/methanol solution for 30 minutes. After washing with phosphate-buffered

saline, cells were blocked before addition of primary antibodies. After blocking

has occurred, CMs were double stained with combinations of the following

primary antibodies: anti-α-actinin, anti-vinculin (Upstate Biotechnology, Lake

Placid, NY), rabbit anti-TRPA1 and/or mouse anti-TRPV1. Alexa Fluor 488- conjugated donkey anti-rabbit and Alexa Fluor 568-conjugated donkey anti-

mouse (Life Technologies) were used as secondary antibodies. Negative

controls included CMs incubated with a single primary antibody and both

secondary antibodies, as well as CMs incubated with secondary antibodies

alone. Images were acquired using an Olympus Fluoview 100 confocal laser

scanning microscope with an X63 objective lens. All images were acquired utilizing a 60X objective with a gain setting of 500 volts and laser excitation wavelengths of 488 nm (multi-line Ar, 8.0% full power, BA505-525 filter) and 543 nm (HeNe, 12.0% full power, BA560-660 filter).

Immunohistochemistry

Hearts were rapidly excised and placed immediately in 4% paraformaldehyde (PFA) and incubated overnight at 4°C. Samples were rinsed and then incubated in 4°C overnight in increasing concentrations of sucrose (10,

15, 30%). Hearts were then frozen into Tissue-Tek OCT tissue-freezing medium

(Sakura Finetek) and prepared for sectioning. Transverse cryosections were taken at 30 µm using a Leica cryostat and mounted on super frost plus coated glass slides. The slides were blocked and stained as previously described

(Russell et al., 2006) using the same antibodies that were applied in the

immunocytochemistry experiments, including DAPI. Negative controls included

TRPA1-/- or TRPV1-/- heart sections stained with TRPA1 and TRPV1 antibodies, respectively, and WT heart sections incubated with secondary antibodies in the absence of a primary. Slides were visualized using a laser scanning confocal

microscope with an oil immersion lens at 10X and 60X. All images were acquired

with a gain setting of 500 volts and laser excitation wavelengths of 488 nm (8.0%

full power, BA505-525 filter) and 543 nm (12.0% full power, BA560-660 filter). Z- stack images were taken at several levels of cardiac myofibers to assess the physical depth of immunostaining and to confirm the localization of TRPA1 and

TRPV1 ion channels at the costameres and Z-discs.

2+ [Ca ]i Measurements

2+ [Ca ]i measurements were performed as previously described (Kanaya et al., 2003). For real-time intracellular calcium measurements, CMs were allowed to adhere to laminin-coated cover slips and incubated at room temperature for 30 min with fura-2 acetoxy methylester (fura-2/AM; 2 µM) in HEPES-buffered saline

(in mM: 118 NaCl, 4.8 KCl, 1.23 CaCl2, 0.6 KH2PO4, 4.6 NaHCO3, 0.6 NaH2PO4,

5.5 glucose, pH 7.4). Coverslips containing the fura-2-loaded CMs were then

mounted on the stage of an Olympus IX-81 inverted fluorescence microscope

(Olympus America). CMs were superfused continuously with HEPES-buffered saline at a flow rate of 2 mL/min and compounds (agonists, antagonists) were delivered for ~10 sec with a 10 min wash period between subsequent treatments.

Potassium chloride (KCl; 35 mM) was utilized to confirm that the CMs are

polarized and respond to KCl-triggered depolarization leading to the activation of

2+ 2+ the voltage-gated L-type Ca channels. [Ca ]I measurements were simultaneously recorded on individual cells using the fluorescence imaging system and Easy Ratio Pro software (Photon Technology International) equipped with a multiwavelength spectrofluorometer (Deltascan RFK6002) and a

QuantEM 512SC electron multiplying camera (Photometrics). Images and real- time calcium tracing data were acquired using an alternating excitation wavelength protocol (340, 380 nm/20 Hz) and emission wavelength of 510 nm.

Background fluorescence was automatically corrected for the experiments using

Easy Ratio Pro. The ratio of the two intensities was used to measure changes in

2+ [Ca ]I due to the fact that calibration of the system relies upon a number of

assumptions. Dose response curves to agonists alone or agonist in the presence

of increasing concentrations of antagonists were performed utilizing a fluorescent

calcium assay kit (Molecular Probes) and a dual-wavelength multimode detector

(DTX 880, Beckamn Coulter) with excitation and emission wavelengths set at

494 and 516 nm, respectively. CMs were centrifuged and resuspended in assay buffer (1X HBSS, 20 mM HEPES) and pipetted into a 96-well plate and then incubated at 37°C for ~1 h to allow the cells to settle. A dye mix (containing Fluo-

4 NW and probenecid) was subsequently added to the wells and incubated for 30 min at 37°C and then for an additional 30 min at room temperature. Increasing concentrations of AITC or capsaicin were aliquoted into each of the wells (10 min prior to measurement) to obtain dose-response curves for agonist-induced

2+ increases in [Ca ]i. Similarly, dose response curves following a 10 min pre-

treatment of increasing concentrations of the antagonists prior to addition of a

maximal dose of channel agoinst were also performed. Results were quantified

and are expressed as mean ± SEM.

Statistical Analysis

2+ [Ca ]I imaging experimental protocols were repeated with a minimum of

six separate coverslips containing CMs from the respective groups. Results from

each coverslip were averaged so each coverslip of CMs were equally weighted in

calculations. The Shapiro-Wilk normality test was used to examine the Gaussian

distribution. Comparisons between the groups were made utilizing repeated-

measures one-way ANOVA and Bonferroni post hoc text (p < 0.05). All results are expressed as mean ± SEM. Error bars in the figures signify the variability of

2+ peak [Ca ]I intensities in calcium imaging experiments or in the calcium assay as

noted in the legends. Statistical analysis was carried out using Sigma Plot 11.0

software (Systat Software).

RESULTS

TRPA1 and TRPV1 are Expressed in CMs

Representative immunoblots demonstrating the expression of TRPA1 and

TRPV1 in CMs are shown in Figures 6A-B. CMs obtained from WT, TRPA1-/- and TRPV1-/- mice were lysed and prepared for immunoblotting using antibodies recognizing TRPA1 or TRPV1. Non-transfected F-11 cells and F-11 cells transfected with TRPA1 and TRPV1 served as the negative and positive controls, respectively. Immunoblot analysis demonstrated expression of TRPA1 at 110 kDa (Figure 6A) and subsequent reprobing of TRPV1 at 95 kDa (Figure 6B).

Figure 6: TRPA1 and TRPV1 are expressed in CMs obtained from wild-type

(WT) mice. Representative immunoblots depicting TRPA1 and TRPV1

expression in adult mouse ventricular cardiomyocytes (CMs). Representative immunoblots demonstrating TRPA1 (A) and TRPV1 (B) expression in F-11 cells transfected with TRPA1 and TRPV1 (F-11 A1/V1), non-transfected F-11 cells (F-

11 NT), wild-type (WT) CMs, TRPA1-/- CMs and TRPV1-/- CMs. GAPDH was

probed as the loading control. PL = protein ladder. n = cells obtained from 6

hearts.

TRPA1 and TRPV1 Colocalize in Cardiac Tissue

Mid-ventricular heart sections obtained from WT, TRPA1-/- and TRPV1-/-

mice were subjected to anti-TRPA1 and/or anti-TRPV1 antibodies and prepared for immunohistochemical analysis (Figure 7). DAPI was used to label nuclei.

Immunohistochemical staining of TRPA1 or TRPV1 in sections obtained from

WT, TRPA1-/- and TRPV1-/- mice are shown in Figure 7A-B and illustrate the presence of TRPA1 and TRPV1 throughout the heart in sections obtained from

WT mice whereas no immunodetectable staining was evident for TRPA1 or

TRPV1 in the sections obtained from TRPA1-/- or TRPV1-/- mice respectively. WT

heart sections were also exposed to secondary antibodies in the absence of

TRPA1 and TRPV1 primary antibodies, which yielded no immunodetectable

labelling (Figure 7C). Mid-ventricular heart sections obtained from WT mice that were incubated with both anti-TRPA1 and anti-TRPV1 antibodies are also illustrated in Figure 7D indicating colocalization of TRPA1 and TRPV1 throughout the endocardium, mid-myocardium and epicardium. Upon further examination through acquisition of serial confocal Z-stack images of the sections to assess physical depth of tissue staining we observed that both TRPA1 and

TRPV1 appear to be localized at the costameres and the Z-discs (Figure 8A-B).

Figure 7: TRPA1 and TRPV1 colocalize throughout the different layers of cardiac

muscle. Representative confocal images (10X magnification) obtained from WT

mouse hearts depicting immunolocalization of TRPA and TRPV1 in the

endocardium (Endo), myocardium (Myo) and epicardium (Epi). 30 μm sections were labeled with antibody recognizing TRPA1 (green; A) or TRPV1 (red; B) in hearts obtained from WT, TRPA1-/- and TRPV1-/- mice. WT heart sections were also treated with secondary antibody in the absence of primary (C). DAPI was used for nuclear staining. Representative confocal images of double-stained sections indicate that immunodetectable TRPA1 and TRPV1 colocalize throughout cardiac tissue (D). n = sections obtained from 6 hearts.

Figure 8: TRPA1 and TRPV1 localize at the costameres and Z-discs in cardiac myofibers. Representative confocal Z-stack images obtained from WT hearts to assess the physical depth of tissue staining reveals that TRPA1 and TRPV1 localize at the costameres (yellow arrows) and Z-discs (white arrows) within the

46 tube-like structure of cardiac myofibers. Heart sections (30 μm) were labeled with antibody recognizing TRPA1 (green; A) or TRPV1 (red; B) and images were acquired at the top and middle (Mid) layers of the myofiber. The lack of Z-disc staining at the middle-levels indicate TRPA1 and TRPV1 predominantly localize toward the outer Z-discs and associated costameric complexes in cardiac tissue.

Scale bar, 10 μm. n = sections obtained from 6 hearts.

Finally, immunocytochemical assessment of the precise cytoskeletal

localization of TRPA1 and TRPV1 was performed in freshly isolated CMs. In

these studies, CMs were immunostained with anti-α-actinin (Z-disc marker), anti- vinculin (costamere and intercalated disc marker), anti-TRPA1 and/or anti-

TRPV1 (Figure 9). Figures 9A-B demonstrates colocalization of both TRPA1 and TRPV1 with α-actinin (panel A) indicating localization of both channels at the z-disc in addition to both channels also colocalizing with vinculin (panel B) indicating their presence at the costameres and intercalated disc. Confirmation of the coexpression of TRPA1 and TRPV1 channels in CMs and their colocalization at the z-discs, costameres and intercalated discs is demonstrated in

Figure 9C.

Figure 9: TRPA1 and TRPV1 colocalize at the Z-disc, costameres and

intercalated discs in CMs. Confocal images in CM obtained from WT mice

confirm that TRPA1 and TRPV1 colocalize at the Z-disc, costameres as well as the intercalated discs. Freshly isolated CMs were double labeled with antibody recognizing TRPA1 or TRPV1 and α-actinin (z-disc marker; white arrows) A).

Similar immunolabeling with antibody recognizing TRPA1 or TRPV1 and vinculin

(costamere and intercalated disc marker, yellow arrow) was also performed (B).

Representative confocal images of double-labeled CMs revealed that TRPA1 and TRPV1 colocalize at the Z-disc, costamere, and intercalated discs (C). Scale bar, 10 μm. n = cells obtained from 6 hearts.

TRPA1 and TRPV1 Agonists Elicit Dose-Dependent Transient Rises in

2+ [Ca ]I in CMs

To examine the extent to which TRPA1 and TRPV1 channels are

physiologically functional in the heart, we performed dose-response studies

2+ assessing changes in [Ca ]i in response to the TRPA1 agonist, AITC or the

TRPV1 agonist, capsaicin in freshly isolated CMs (Figures 10 and 11). Real-

2+ time calcium measurements revealed transient rises in [Ca ]I in WT CMs when

treated with AITC (100 μM; Figure 10A) or capsaicin (100 nM; Figure 10B). The

2+ AITC- and capsaicin-induced transient rises in [Ca ]i were absent in CMs

obtained from TRPA1-/- or TRPV1-/-, respectively (Figure 10C-D). CMs from all three groups of mice were exposed to potassium chloride (KCl; 35 mM) subsequent to agonist activation to confirm cell viability and adequate fura-2AM loading. -Similarly, pharmacological studies indicate that the transient rises in

2+ [Ca ]i elicited via AITC and capsaicin were eliminated in WT CMs pretreated

with TRPA1 antagonist, HC-030031 (500 nM; Figure 10E) or the TRPV1

antagonist SB366791 (10 μM; Figure 10F), respectively. Moreover, capsaicin

2+ -/- and AITC induced transient rises in [Ca ]I in CMs obtained from TRPA1 and

TRPV1-/- mice, respectively (Figure 10G & 10H). Summarized data illustrating

2+ the effects of AITC or capsaicin on transient rises in [Ca ]i in CMs pretreated

with pharmacological inhibitors of the channels or in CMs obtained from TRPA1-/-

and TRPV1-/- mice are depicted in Figure 10G.

Figure 10: TRPA1 and TRPV1 stimulation elicits transient increases in intracellular free calcium concentration in quiescent CMs. Representative traces

depicting the effect of TRPA1 agonist stimulation with AITC (100 μM; 10 second

exposure) or TRPV1 agonist stimulation with capsaicin (100 nM; 10 second

2+ exposure) on [Ca ]i in freshly isolated CMs obtained from WT mice (A and B,

respectively) as well as in CM’s obtained from TRPA1-/- or TRPV1-/- mice (C and

D, respectively). Representative traces depicting the effect pre-treatment with the TRPA1 antagonist, AITC HC-030031 (500 nM) or the TRPV1 antagonist,

2+ SB366791 (10 μM) on AITC- or capsaicin-induced transient rises in [Ca ]i in

CMs obtained from WT mice (E and F, respectively). Representative traces

depicting the effect of AITC or capsaicin on CMs obtained from TRPA1-/- and

TRPV1-/- mice (G and H, respectively). CMs were treated with potassium chloride (KCl) where indicated. Summarized data for A-H (I). Data are expressed as a percent of the response observed in vehicle-treated CMs (% of control mean value ± SEM). n = experiments performed in CMs obtained from 6 separate mice.

*P < 0.05 compared to vehicle-treated cells (ethanol). #P < 0.05 compared to

AITC-treated WT CMs. †P < 0.05 compared to capsaicin-treated WT

CMs. Statistical analysis performed using one way analysis of variance and the

Bonferroni post hoc test. n = cells obtained from 6 hearts.

Finally, we assessed the dose-dependency of AITC and capsaicin to elicit

2+ increases in [Ca ]i as well as the dose-dependent effects of the specific TRPA1

and TRPV1 antagonists to block the responses to AITC or capsaicin in CMs

(Figure 11). These experiments were performed using a 96 well plate fluorescent

Ca2+ assay kit where cells in each well could be stimulated with only one dose of the agonist and/or inhibitor prior to the measurement. This was done in order to circumvent the potential for repetitive stimulation of the TRPA1 or TRPV1 channels to desensitize. Summarized data depicting the dose-dependent effect

2+ of AITC or capsaicin on [Ca ]i in CM obtained from WT and TRPA1-/- or TRPV1-

/- mice are depicted in Figure 11A-B. Summarized data depicting the dose- dependent effect of the TRPA1 antagonist, HC-030031 or the TRPV1 antagonist,

2+ SB366791 on AITC-induced or capsaicin-induced increases in [Ca ]i,

respectively, are depicted in Figure 11C-D.

Figure 11: AITC and capsaicin induce dose-dependent increases in intracellular

free calcium concentration in WT CMs through mechanisms dependent upon

TRPA1 and TRPV1, respectively. Summarized data depicting the dose-

2+ dependent effect of AITC or capsaicin on [Ca ]i in CMs obtained from WT,

TRPA1-/- or TRPV1-/- mice (A and B respectively). Responses to ethanol alone

(vehicle control) were normalized to 100% and considered the control response.

Summarized data depicting the dose-dependent effect of the TRPA1 antagonist,

HC-030031 or the TRPV1 antagonist, SB366791 on AITC- (100 µM) or

2+ capsaicin- (100 nM) induced increases in [Ca ]i. (C and D respectively).

Responses to the AITC alone (100 µM) were normalized to 100% and considered the control response. n = experiments performed in CMs obtained from 6 separate mice. *P<0.05 compared to vehicle treated (ethanol) control.

Statistical analysis performed using one way analysis of variance and the

Bonferroni post hoc test. n = cells obtained from 6 hearts.

DISCUSSION

To our knowledge, this is the first study to thoroughly characterize the

ultrastructural localization and functional expression profiles of TRPA1 and

TRPV1 ion channels in adult mouse CMs. The immunodetection, ultrastructural

localization and functionality of TRPA1 channels at the protein level in cardiac

muscle has not been previously reported. Although immunodetectable TRPV1

has previously been identified in mouse hearts (Gao et al., 2014; Zhong and

Wang, 2009) and appears to be located on the epicardial surface as well as in blood vessels and perivascular nerves (Zhong and Wang, 2009), the precise ultrastructural location of TRPV1 channels in the hearts as well as a detailed pharmacological profile of the channel has yet to be established in adult mouse

CMs.

The major findings of the current study are that both TRPA1 and TRPV1 are co-expressed in the adult mouse heart throughout the epicardium, myocardium as well as endocardium, and both channels appear to co-localize at the costameres, z-disc and intercalated discs in isolated CMs. Moreover, both

TRPA1 and TRPV1 channels are functional in isolated CMs since both channels

2+ respond to selective agonist stimulation with a transient increase in [Ca ]i in a

dose-dependent manner, an effect that is dose-dependently attenuated with

specific channel antagonists and is absent all together in CMs obtained from

TRPA1-/- and TRPV1-/- mice.

TRPA1 and TRPV1 Expression in Cardiac Tissue

The superfamily of TRP ion channels play important roles in the physiology of the cardiovascular system by regulating fundamental cell functions such as contraction, relaxation, proliferation, differentiation and cell death (Yue et

al., 2015), but also play an important role in the pathophysiology of many

diseases in the cardiovascular system (Inoue et al., 2006; Vennekens, 2011;

Watanabe et al., 2008). Although the expression, ultrastructural localization and

physiological/pathophysiological role(s) for TRPA1 ion channels in myocardial

tissue has yet to be determined, the expression of TRPV1 channels in cardiac

muscle and their role in physiological/pathophysiological processes in the heart

have recently been reported and are rapidly emerging as key players in a myriad

of cellular and molecular events related to cardiac diseases. For example,

several studies have demonstrated the presence of TRPV1 in cardiac muscle

and an important role for the channel in mediating myocardial protection from

ischemic injury (Huang et al., 2009; Sexton et al., 2007; Wang and Wang, 2005;

Zhong and Wang, 2007, 2009). In contrast, evidence also exists for TRPV1 channels playing a role in the development of the pathophysiology of cardiac hypertrophy and heart failure (Buckley and Stokes, 2011; Horton et al., 2013)

whereas another study indicated TRPV1 activation attenuates high-salt diet- induced cardiac hypertrophy and fibrosis (Lang et al., 2014). Limited information is available describing where in the heart TRPV1 channels are expressed, which specific cell types are involved (myocytes, fibroblasts, vasculature, etc) and where is the precise ultrastructural location of the channel in cells where it is

expressed. Previous studies by our laboratory have identified that the sensitivity

of TRPV1 channels to agonist activation can be modulated by TRPA1 channel

agonists, indicating cross-talk/regulation between the channels that may be vital for altering cellular responses to noxious stimuli in sensory neurons (Sinharoy et al., 2015; Wickley et al., 2010; Zhang et al., 2011). Because TRPA1 protein has not yet been reported in cardiac muscle, but plays an important regulatory role in sensory neurons when co-expressed with TRPV1, our objectives were to identify the extent to which TRPA1 is co-expressed with TRPV1 in cardiac muscle and determine their precise ultrastructural locations within the CM. Identifying the localization of these channels could provide important fundamental insight and clarification into their roles in physiological and pathophysiological consequences.

Our studies clearly indicate that both TRPA1 and TRPV1 are expressed throughout the heart and appear to localize at the costameres, z-discs and intercalated discs in adult mouse CMs. Costameres are critically important cytoskeletal structures demonstrated to have significant roles in mechanosensation and bidirectional signal transduction in CMs (Jaka et al.,

2015; Knoll, 2015). Functionally, they are similar to focal adhesion complexes but

they are found in register with Z-discs and circumferentially couple the cardiac myofibrils to the sarcolemma. The positioning of costameres within the ultrastructure of the CM alludes to their vast ability to modulate physiological and pathophysiological events in cardiac tissue. Furthermore, Z-discs are commonly regarded as “internal costameres”, by which they share similar functional roles in

CM biochemical signal transduction; each are implicated in sensing mechanical

stress and subsequent conversion of stress signals into alterations of protein

synthesis, cell-cell communication, protein assembly within the sarcomere and

ion channel function, among others (Samarel, 2015). The exact mechanisms by

which mechanical stimuli are converted into biochemical responses remain

elusive, however, mechanosensitive ion channels have been implicated as

potential players by which they would mediate mechanisms that occur between

the cytoskeleton and the sarcolemma (Ervasti, 2003). Although the roles that

TRPA1 and TRPV1 channels play in mediating intracellular signaling pathways

and molecular mechanisms within cardiac tissue remains elusive, these channels

are understood to be mechanosensitive signal transducers in other cell types throughout the body (Fernandes et al., 2011; Inoue et al., 2009) – characteristics

which may allude to which intracellular and intercellular events the channels play

a role in regulating within CMs. Since TRPA1 and TRPV1 have been

demonstrated to serve roles in mechanosensation in tissues throughout the body

(Brierley et al., 2011; Lin et al., 2015; Sappington et al., 2015), we speculate they

will serve a similar role in ventricular CMs, by which they can sense external

stimuli and transmit that signal internally. Furthermore, the localization of TRPA1

and TRPV1 at the region of the presumed intercalated discs suggests the ion

channels may serve a role in cell-cell adhesion or the propagation of chemical

and electrical signals through the network of the lattice-like cardiac tissue.

Even though costameres are generally investigated in regards to their

ability to transmit contractile forces from the inside of CMs to neighboring tissues,

60 as well as converting forces generated in the extracellular matrix to intracellular biochemical signals, costameres and Z-discs are also known to be muscle cell signaling “hot-spots” (Samarel, 2015). Although there is a paucity of information underlying the signaling pathways elicited via TRPA1 and/or TRPV1 activation in

CMs, the ion channels have been demonstrated to induce a myriad of intracellular signaling events upon stimulation in other cell types (Lin et al., 2015;

Sinharoy et al., 2015; Zhang et al., 2011). Indeed, mechanistically delineating intracellular signaling events elicited via TRPA1 and TRPV1 stimulation and the potential role for cross-regulation of the channels in mediating cellular events in cardiac muscle, particularly in mediating myocardial protection, is of great clinical significance and will be the focus of future investigations.

Regulation of cardiac tissue by TRP channels is typically attributed to

TRP-channel-expressing cardiac nerve afferents innervating the heart tissue, itself (Pan and Chen, 2004; Zvara et al., 2006). Consequentially, TRP channel physiological and pathophysiological functionality within cardiac tissue/CMs has yet to be fully elucidated. As noted earlier, a previous study indicated that TRPV1 appeared to localize predominantly to the epicardial layer of the heart (Zhong and Wang, 2009); however, results included within Chapter two of this dissertation indicate that both TRPA1 and TRPV1 are expressed throughout the three layers (endocardium, myocardium, epicardium) of cardiac tissue. Although previous investigations have demonstrated the role of TRPV1 in hypertrophy and heart failure (Buckley and Stokes, 2011; Horton et al., 2013), the extent to which

TRPA1 and TRPV1 regulate basal cardiac function remains elusive.

Furthermore, although TRPV1 has been extensively investigated in regards to its

role in myocardial protection following ischemia (Huang et al., 2009; Sexton et

al., 2007; Wang and Wang, 2005; Zhong and Wang, 2007, 2009), further studies

are required in order to precisely determine how TRPA1 and TRPV1 channel

function is altered in pathological conditions.

TRPA1 and TRPV1 Stimulation Elicits Calcium Influx in CMs

As stated previously, TRPA1 and TRPV1 are non-selective cation

channels that tend to show high permeability to calcium (Fernandes et al., 2011).

Although we demonstrate co-expression and co-localization of both TRPA1 and

TRPV1 in CMs, we next sought to determine whether the channels are functional

and explore their sensitivity to agonist activation which should result in transient

2+ increases in [Ca ]i in CMs. Our studies indicate that the selective agonists for

TRPA1 (AITC) or TRPV1 (capsaicin) channels causes a dose-dependent

2+ transient rise in [Ca ]i. Moreover, this effect was absent in CMs obtained from

TRPA1-/- and TRPV1-/- mice indicating that AITC and capsaicin selectively

stimulate calcium influx through functional TRPA1 and TRPV1, respectively.

2+ Similarly, the agonist-induced transient rise in [Ca ]i was dose-dependently attenuated when CMs obtained from WT mice were pretreated with selective antagonists of TRPA1 (HC-030031) or TRPV1 (SB366791) prior to agonist

2+ stimulation. These transient increases in [Ca ]i are similar to our previous

findings in sensory neurons and indicate functional gating of the channels in

response to the agonists as well as pharmacological sensitivity to well

2+ established antagonists of each channel. A dose-dependent increase in [Ca ]i was previously observed in a cultured cardiac cell line (H9c2 cells), however the

2+ response to capsaicin was a sustained elevation in [Ca ]i which did not return to baseline, suggesting that the TRPV1 channels in this cell line may not be gating properly (Gao et al., 2014).

Calcium regulation in CMs partially dictates inotropic, chronotropic and lusitropic properties and the resulting cellular energetics within cardiac tissue.

Although TRP channel-mediated calcium entry has been shown to induce intracellular signaling cascades in several other cell types (Ambudkar, 2016;

Kurosaka et al., 2016; Stueber et al., 2016), as well as induce fibroblast proliferation and differentiation leading to various forms of arrhythmia, hypertrophy or heart failure (Davis et al., 2012; Du et al., 2010; Harada et al.,

2012), the extent to which TRPA1 and TRPV1 modulate these effects in CMs remains to be determined and will be the focus of future investigations. Taken together with the polymodal activation characteristics of TRP channels, we speculate that TRPA1 and TRPV1 may have diverse functions in cardiac physiology and pathophysiology. However, since TRP channels are able to integrate and initiate signaling events via calcium entry and consequential membrane depolarization, it is feasible to hypothesize a role for these channels in mediating cellular functions such as contraction, relaxation, myogenic regulation (perhaps due to mechanosensation) and cell death in the heart.

Summary and Conclusions of Chapter 2

Due to the paucity of information regarding the functional expression and

localization of TRPA1 and TRPV1 in cardiac tissue, this investigation was

designed to examine expression patterns of the ion channels in freshly isolated

CMs and to begin delineating the physiological functions of the channels in

response to agonist stimulation. Overall, our current findings are consistent with

previous reports of TRPV1 expression in the heart (Pei et al., 2014; Yue et al.,

2015; Zhong and Wang, 2009), but the reporting of TRPA1 at the protein level in

CMs is novel in nature. Furthermore, we demonstrate that TRPA1 and TRPV1

are expressed throughout the different layers of the heart and they colocalize at

the intercalated discs, but are most heavily concentrated at the Z-discs and

costameres within the CM cytoskeleton. The localization of TRPA1 and TRPV1 in

CMs have prompted several hypotheses. First, the Z-disc is the site of

localization for many proteins, which indicate that the ion channels may share similar signaling pathways and/or are involved in direct physical interactions with other structures located therein. Secondly, CMs have stress-strain sensors embedded at several locations, including the Z-disc, costameres and intercalated discs; this suggests a potential role for the receptors in mediating mechanotransduction (Hoshijima, 2006). Lastly, the localization of the channels at the intercalated discs could be correlated with the presence of proteins which mediate calcium-dependent cell-to-cell adhesion, such as N-cadherin (Li, 2014;

Sheikh et al., 2009). Moreover, stimulation of TRPA1 and TRPV1 in freshly

2+ isolated CMs induces dose-dependent, transient rises in [Ca ]I – effects which

64 can be dose-dependently eliminated through the use of selective channel antagonists. In order to identify the myriad of events likely to be modulated by the presence and activation of TRPA1 and TRPV1 in cardiac tissue, future studies are required to further elucidate whether the channels communicate (either directly or indirectly) and the extent to which they mediate physiological events.

In conclusion, the results included herein provide a foundation for future investigations designed to determine the precise physiological functions of

TRPA1 and TRPV1 in cardiac tissue. Delineating the signal transduction pathways and molecular mechanisms to which they are involved will provide fundamental insight into uncovering novel information regarding the regulation of

TRPA1- and TRPV1-mediated physiological and pathophysiological events in cardiac tissue.

CHAPTER THREE

Stimulation of TRPA1 and TRPV1 Ion Channels Increase Intracellular Ca2+

Transients and Contraction in Mouse Ventricular Myocytes

INTRODUCTION

The transient receptor potential (TRP) ion channel of the ankyrin 1

(TRPA1) and vanilloid 1 (TRPV1) subtypes are members of the TRP superfamily

of structurally related channels. Similar to other TRP ion channel superfamily

members, TRPA1 and TRPV1 act as non-selective cation channels that are

highly permeable to calcium (Fernandes et al., 2011). The TRPA1 ion channel

was first discovered in human fetal lung fibroblasts (Jaquemar et al., 1999) and

later found to be primarily expressed in sensory neurons of the dorsal root,

nodose and trigeminal ganglion (Nagata et al., 2005; Story et al., 2003) where it

is co-expressed with TRP ion channels of the vanilloid receptor subtype 1

(TRPV1) and together, both were proposed to serve a role in pain by acting as

transducers of noxius stimuli (Story et al., 2003). However, over the past decade it has become increasingly evident that TRPA1 and TRPV1 ion channels are

expressed in a variety of tissues and cell types including those of the

cardiovascular system (Nilius, 2007; Vennekens, 2011; Yue et al., 2015).

The findings from Chapter two demonstrate the expression of TRPA1 at

the protein level in cardiac muscle and its co-localization with TRPV1 ion

channels at the costamere, z-disc and intercalated discs in murine

cardiomyocytes (CM). We also demonstrated in the same studies that both

TRPA1 and TRPV1 ion channels were functional and responded to agonist

stimulation by allyl isothiocyanate (AITC) and capsaicin, respectively, with a

2+ 2+ predicted transient rise in intracellular free Ca concentration ([Ca ]i). However, the extent to which TRPA1 or TRPV1 ion channel stimulation plays a role in the modulation or regulation of CM contractile function has not been previously reported. In the current study, we tested the hypothesis that stimulation of

TRPA1 and TRPV1 ion channels in electrically-stimulated mouse ventricular CMs

2+ results in increases in [Ca ]i dynamics and a concomitant increase in CM contractile function. We found that stimulation of both TRPA1 and TRPV1 result

2+ in dose-dependent increases in peak [Ca ]i and a concomitant increases in CM fractional shortening. We also demonstrate a dose-dependent acceleration in CM

2+ time to peak [Ca ]i and velocity of shortening as well as an acceleration in the

2+ 2+ CM [Ca ]i decay and the velocity of relengthening. The effects of AITC on [Ca ]i

and contractile function were not observed in CMs pretreated with the TRPA1

antagonist, HC-030031, nor in CMs obtained from TRPA1 null (TRPA1-/-) mice.

2+ Similarly, the effects of capsaicin on [Ca ]i and CM contractility were not

observed in CMs pretreated with the TRPV1 antagonist, SB366791 nor in CMs

67 obtained from TRPV1 null (TRPV1-/-) mice. In vivo preparations where AITC was administered intravenously yielded similar results as evidenced by a dose- dependent increase in ejection fraction in mouse hearts. These data indicate that stimulation of TRPA1 and TRPV1 ion channels in cardiac muscle results in activation of cellular signal transduction pathways associated with increasing

2+ [Ca ]i and contractile function in adult mouse ventricular CMs. TRPA1- and

TRPV1-dependent signaling pathways may represent a novel mechanism for increasing the inotropic and lusitropic state of the heart.

MATERIALS AND METHODS

Animal Model

Twelve 4-month-old male C57BL/6 mice (n = 6/group), four TRPA1-/- male mice and three TRPV1-/- male mice (Jackson Labs, Bar Harbor, ME) were used and maintained in accordance with the Guide for the Care and Use of Laboratory

Animals (NIH). All animals were housed at the Kent State University animal care facility (Kent, OH), which is accredited by the American Association for

Accreditation of Laboratory Animal Care.

Isolation of CMs

Murine hearts were excised and transferred to a Langendorff apparatus for CM isolation, as previously described. In brief, mice were sacrificed via cervical dislocation and hearts were rapidly excised then placed into a dish

containing perfusion buffer. After the aorta was cannulated and blood was

flushed, the hearts were subjected to retrograde perfusion at 37°C and pH 7.4

with a modified Krebs-Henseleit buffer (in mM: 120.4 NaCl, 4.8 KCl, 0.6 KH2PO4,

0.6 Na2HPO4, 1.2 MgSO4-7HsO, 10 Na-HEPES, 4.6 NaHCO3, 30 taurine, 10

BDM, and 5.5 glucose). The calcium-free buffer was sterile-filtered and paced

with a peristaltic pump (Masterflex) to begin retrograde perfusion of the heart at a

rate of 4 mL/min. After perfusion for 4 minutes, the same solution containing

collagenase type II (300 U/mg, Worthington Biochemical) perfused the heart for

an additional 8 minutes until the heart became soft. The left ventricles were

removed, minced, then triturated in Krebs-Henseleit buffer containing fetal bovine serum. The resulting cellular digest was washed and resuspended in HEPES- buffered saline (in mM: 118 NaCl, 4.8 KCl, 0.6 KH2PO4, 4.6 NaHCO3, 0.6

NaH2PO4, 5.5 glucose, pH 7.4) at 23°C. CM yield was typically ~80-90%. CMs

were then either subjected to slow calcium reintroduction ([1.23 mM]) and

2+ subsequent [Ca ]i and shortening protocols.

2+ Simultaneous Measurement of [Ca ]i and Shortening

2+ Simultaneous measurement of [Ca ]i and contractile function was

performed in individual freshly isolated CMs as previously described by our

laboratory (Kurokawa et al., 2002). CMs were incubated at room temperature for

30 minutes with fura-2 acetoxy methylester (fura-2/AM; 2 µM) in HEPES-buffered saline (in mM: 118 NaCl, 4.8 KCl, 1.23 CaCl2, 0.8 MgSO4-7H2O, 0.6 KH2PO4, 4.6

NaHCO3, 0.6 NaH2PO4, 5.5 glucose, pH 7.4). Coverslips containing the fura-2-

loaded CMs were then mounted on the stage of an Olympus IX-71 inverted

fluorescence microscope (Olympus America). CMs were superfused

continuously with HEPES-buffered saline at a flow rate of 2 mL/min and compounds were introduced in a dose-dependent manner. Sarcomere

2+ shortening and [Ca ]I measurements were simultaneously recorded on individual

cells using the fluorescence imaging system and Easy Ratio Pro software

(Photon Technology International) equipped with a multiwavelength

spectrofluorometer (Deltascan RFK6002) and a QuantEM 512SC electron

multiplying camera (Photometrics). Images and real-time calcium tracing data were acquired using an alternating excitation wavelength protocol (340, 380 nm/20 Hz) and emission wavelength of 510 nm. Background fluorescence was automatically corrected for the experiments using Easy Ratio Pro. The ratio of

2+ the two intensities was used to measure changes in [Ca ]I due to the fact that

calibration of the system relies upon a number of assumptions. Hardware and

software for data acquisition and analysis were generously provided by Horiba

Scientific (Edison, NJ).

2+ Analysis of [Ca ]i and Shortening Data

The following variables were calculated for each individual contraction:

sarcomere length (µm), fractional shortening (% of sarcomere length change

during shortening), maximum velocity of cell shortening and relengthening

2+ 2+ 2+ (µm/sec), peak [Ca ]i (340/380 ratio), time to peak [Ca ]i (msec), [Ca ]i decay

2+ to baseline (msec), slope of time to peak [Ca ]i, time to 50 and 90%

2+ relengthening (msec) and time to 50 and 90% [Ca ]i decay to baseline (msec).

Variables from 10 contractions were averaged to obtain mean values at baseline and in response to the intervention. Averaging the variables over time minimizes beat-to-beat variation.

Transthoracic Echocardiography

In vivo cardiac functional parameters were assessed utilizing a Vevo 770

(VisualSonics, Inc., Toronto, Ontario, Canada) in the following animal groups: wild-type mice with acute AITC infusion via jugular catheterization. Mice were anesthetized using 2% isoflurane and placed on an adjustable platform equipped with ECG electrodes to monitor heart and respiration rates. Doppler echocardiography was carried out parasternally across a shaved chest wall and short (midpapillary level)- and long-axis images were obtained. Functional measurements were averaged from three cardiac cycles. The following parameters were assessed as indicators of cardiac function: LV diastolic internal diameter (LVIDd), LV systolic internal diameter (LVIDs), posterior wall thickness and ejection fraction (LVIDd-LVIDs/LVIDs × 100). Measurements were calculated blinded reviewers using the Vevo 770/3.0 software.

Statistical Analysis

Dose response curves to AITC, capsaicin and isoproterenol were repeated in CMs obtained from 6 different wild-type mouse hearts. The effects of

AITC and capsaicin on TRPA1-/- and TRPV1-/- CMs were conducted in four and three different hearts, respectively. The effects of HC030031 and SB366791 were repeated in CMs obtained from 3 wild-type mouse hearts. Results obtained from each heart were averaged so that all hearts were weighted equally. Within group comparisons were made using one-way analysis of variance with repeated measures and the Bonferroni post hoc test. Differences were considered statistically significant at p < 0.05. All results are expressed as means ± SEM.

RESULTS

2+ AITC Stimulates Dose-Dependent Increases in Peak [Ca ]i and Contractile

Function in CMs

Figure 12 depicts representative traces demonstrating a marked increase

2+ in shortening (change in sarcomere length; Figure 12A) and peak [Ca ]i

(change in 340/380 ratio; Figure 12B) in an individual CM following treatment with the TRPA1 agonist, AITC (100 µM). Exploded views illustrating dose-

2+ dependent changes in sarcomere length and [Ca ]i following exposure to AITC

(1-300 µm) are depicted in panels C and D, respectively. Summarized data

2+ demonstrating dose-dependent changes in [Ca ]i dynamics and contractile

function following exposure to AITC are shown in Table I. The AITC-induced (100

2+ µM) increase in [Ca ]i and contractile function were completely blocked by pre-

treatment with the TRPA1 antagonist, HC-030031 (10 µM), resulting in a peak

2+ [Ca ]i and fractional shortening that were 98 ± 3.1% and 101 ± 2.2% of steady

state baseline control value, respectively.

2+ Figure 12: Allyl isothiocyanate (AITC) increases [Ca ]I and shortening in CMs.

Original traces demonstrating the effect of the TRPA! agonist, AITC (100 µM), on

2+ steady state sarcomere length (panel A) and [Ca ]i (panel B) in an individual mouse ventricular myocyte. AITC was added where indicated on the figure.

2+ Changes in sarcomere length were measured in micrometers. [Ca ]i was measured as the 340/380 ratio. Exploded views depicting the dose-dependent

2+ changes in sarcomere length and [Ca ]i (panels C and D, respectively) before

(control) and after addition of AITC (1-300 µM) to single ventricular myocyte.

AITC was added to the bath in a cumulative fashion.

AITC Increases Fractional Shortening, Maximum Velocity of Shortening and

Maximum Velocity of Relengthening in CMs

Figure 13 represents overlays depicting the dose-dependent increases in

CM fractional shortening (Figure 13A), maximum velocity of shortening (Figure

13C) and maximum velocity of relaxation (Figure 13E) following treatment with

AITC (1-300 µM). AITC (100 µM) increased fractional shortening, maximum velocity of shortening and maximum velocity of relaxation to 195 ± 6.8%, 213 ±

7.9% and 196 ± 8.7% of control, respectively. The summarized dose response data for panels A, C and E are depicted in panels B, D and F, respectively and are expressed as a percent of control. The summarized raw data for these parameters are listed in Table I and are expressed as % change in sarcomere length (fractional shortening) and µm/sec (velocity of shortening/relengthening).

Figure 13: AITC increases fractional shortening, maximum velocity of shortening and maximum velocity of relengthening in CMs. Overlays of individual cell shortening and relengthening events illustrating the dose-dependent effects of

AITC (1-300 µM) on fractional shortening are depicted in panel A.

Representative overlays assessing changes in sarcomere length normalized to peak height (set at 100%) and aligned at initiation of stimulus or at peak

shortening to illustrate dose-dependent changes in timing of shortening and relengthening are depicted in panels C and E, respectively. Summarized data for panels A (fractional shortening), C (maximal velocity of shortening) and E

(maximal velocity of relengthening) are depicted in panels B, D and F, respectively. Results are expressed as percent of steady state baseline control

(Ctrl) value set at 100%. Changes in fractional shortening were measured as percent of sarcomere length. Changes in velocity were measured in micrometers/sec. * P < 0.05 compared with Ctrl. n = 18 cells from 6 hearts.

2+ 2+ AITC Increases Peak [Ca ]i and Accelerates Time to Peak [Ca ]i and the

2+ Rate of [Ca ]i Decay in CMs

Figure 14 represents overlays depicting the dose-dependent increases in

2+ 2+ peak [Ca ]i (Figure 14A) as well as an acceleration in time to peak [Ca ]i

2+ (Figure 14C) and in [Ca ]i decay (Figure 14E) following treatment with AITC (1-

2+ 300 µM). AITC (100 µM) increased peak [Ca ]i to 180 ± 14.0% of control. Time

2+ 2+ to peak [Ca ]i and the Ca decay were accelerated by 63 ± 3.1% and 45 ±

2.9%, respectively, compared to control. The summarized data for panels A, C and E are depicted in panels B, D and F, respectively and are expressed as a percent of control. The summarized raw data for these parameters are listed in

Table 1 and are expressed in msec.

2+ 2+ Figure 14: AITC increases peak [Ca ]I and accelerates time to peak [Ca ]I and

2+ the rate of [Ca ]I decay in CMs. Overlays illustrating the dose-dependent effects

2+ of AITC (1-300 µM) on peak [Ca ]i are depicted in panel A. Representative

2+ overlays assessing dose-dependent changes in [Ca ]i peak amplitude normalized to peak height (set at 100%) to illustrate changes in time to peak

2+ 2+ [Ca ]i and the time of [Ca ]i decay are depicted in panels C and E, respectively.

2+ 2+ Summarized data for panels A ([Ca ]i peak amplitude), C (time to peak [Ca ]i)

2+ and E ([Ca ]i decay) are depicted in panels B, D and F, respectively. Results are expressed as percent of steady state baseline control (Ctrl) value set at 100%.

2+ Changes in peak [Ca ]i are measured as the change in the 340/380 ratio.

Changes in timing are measured in milliseconds. Individual traces were

smoothed using the Savitzky-Golay filter to increase the signal-to-noise ratio. * P

< 0.05 compared with Ctrl. n = 18 cells from 6 hearts.

2+ Capsaicin Stimulates Dose-Dependent Increases in [Ca ]i and Contractile

Function in Electrically-Paced CMs

Figure 15 depicts exploded views illustrating dose-dependent changes in

2+ sarcomere length (change in sarcomere length; Figure 15A) and [Ca ]i (change

in 340/380 ratio; Figure 15B) in an individual CM following treatment with

capsaicin (0.1-1 µM). Summarized data depicting the dose-dependent increases

in CM fractional shortening, maximum velocity of shortening and maximum

velocity of relaxation following exposure to capsaicin (0.1-1 µM) are shown in

panels C, E and G, respectively. Capsaicin (0.1 µM) increased fractional

shortening, maximum velocity of shortening and maximum velocity of relaxation

to 197 ± 8.8%, 187 ± 9.7% and 178 ± 12.0% of control, respectively. Summarized

2+ data depicting the dose-dependent increases in peak [Ca ]i as well as an

2+ 2+ acceleration in time to peak [Ca ]i and in [Ca ]i decay following exposure to

capsaicin (0.1-1 µM) are shown in panels D, F and H, respectively. The TRPV1

2+ agonist, capsaicin (0.1 µM), increased peak [Ca ]i to 200 ± 11.1% of control.

2+ 2+ Time to peak [Ca ]i and the Ca decay were accelerated by 53 ± 3.2% and 48

± 1.7%, respectively, compared to control. The summarized data for panels C-H

are expressed as a percent of control. Summarized data demonstrating dose-

2+ dependent changes in [Ca ]I dynamics and contractile function following

exposure to capsaicin are shown in Table II and are expressed as % change in

sarcomere length (fractional shortening), µm/sec (velocity of

2+ 2+ shortening/relengthening) and msec (time to peak [Ca ]I and [Ca ]I decay). The

2+ capsaicin-induced (0.1 µM) increase in [Ca ]i and contractile function were

81 completely blocked by pre-treatment with the TRPV1 antagonist, SB366791 (10

2+ µM), resulting in a peak [Ca ]i and fractional shortening that were 99 ± 1.7% and

102 ± 3.7% of steady state baseline control value, respectively.

2+ Figure 15: Capsaicin increases [Ca ]I and contractile function in CMs.

Representative traces demonstrating the effect of capsaicin (0.1 – 1 µM) on

2+ steady state sarcomere length (panel A) and [Ca ]i (panel B) in an individual

mouse ventricular myocyte. Changes in sarcomere length were measured in

2+ micrometers. [Ca ]i was measured as the 340/380 ratio. Summarized data for

2+ fractional shortening, maximal velocity of shortening/relengthening, [Ca ]i peak

2+ 2+ amplitude, time to peak [Ca ]I and [Ca ]i decay are depicted in panels C-H.

Results are expressed as percent of steady state baseline control (Ctrl) value set

at 100%. Changes in fractional shortening were measured as percent of

sarcomere length. Changes in velocity were measured in micrometers/sec.

2+ Changes in peak [Ca ]i are measured as the change in the 340/380 ratio.

Changes in timing are measured in milliseconds. Individual traces were

smoothed using the Savitzky-Golay filter to increase the signal-to-noise ratio. * P

< 0.05 compared with Ctrl. n = 12 cells from 6 hearts.

2+ AITC Induces Increases in [Ca ]i and Contractile Function via a TRPA1-

Dependent Process in CMs

Figure 16 depicts exploded views of representative traces demonstrating

2+ that AITC (100 µM) has no effect on contractile function (Figure 16A) or [Ca ]i

(Figure 16B) in CMs obtained from TRPA1-/- mice. In CMs obtained from TRPA1-

/- 2+ mice, peak [Ca ]i and fractional shortening were 102 ± 4.1% and 101 ± 5.2% of

steady state baseline control, respectively, following exposure to AITC.

Treatment of CMs obtained from TRPA1-/- mice with capsaicin (0.1 µM)

2+ demonstrated a similar effect on contractile function (Figure 16C) and [Ca ]i

(Figure 16D) to what is observed in CMs obtained from wild-type mouse hearts.

2+ Following treatment with capsaicin (0.1 µM), peak [Ca ]i and fractional

shortening were 198 ± 14.6% and 190 ± 12.1%, respectively, in CMs obtained

from TRPA1-/- mice (summarized data not shown).

2+ Figure 16: AITC has no effect on [Ca ]I and shortening in CMs obtained from

TRPA1 null mice. Exploded views of original traces demonstrating the lack of

2+ µ effect of AITC (100 M) on steady state sarcomere length (panel A) and Ca ]i

(panel B) in an individual mouse ventricular myocyte obtained from TRPA1 null

-/- mice (TRPA1 ). Representative traces demonstrating the effect capsaicin (0.1

2+ µM) on sarcomere length (panel C) and Ca ]i (panel D) in an individual CM

-/- obtained from a TRPA1 mouse. Changes in sarcomere length were measured

2+ in micrometers. [Ca ]i was measured as the 340/380 ratio.

2+ Capsaicin-Induced Increases in [Ca ]i and Contractile Function Occur

Through a TRPV1-Dependent Mechanism in CMs

Figure 17 depicts exploded views of representative traces demonstrating

that capsaicin (0.1 µM) has no effect on contractile function (Figure 17A) or

2+ -/- [Ca ]i (Figure 17B) in CMs obtained from TRPV1 mice. In CMs obtained from

-/- 2+ TRPV1 mice, peak [Ca ]i and fractional shortening were 104 ± 6.3% and 100 ±

1.8% of steady state baseline control, respectively, following exposure to

capsaicin. Treatment of CMs obtained from TRPV1-/- mice with AITC (100 µM)

2+ demonstrated a similar effect on contractile function (Figure 17C) and [Ca ]i

(Figure 17D) to what is observed in CMs obtained from wild-type mouse hearts.

2+ Following treatment with AITC (100 µM), peak [Ca ]i and fractional shortening

were 201 ± 17.1% and 196 ± 13.0%, respectively, in CMs obtained from TRPV1-/-

mice (summarized data not shown).

2+ Figure 17: Capsaicin has no effect on [Ca ]I and shortening in CMs obtained from TRPV1 null mice. Exploded views of original traces demonstrating the lack

of effect of capsaicin (0.1 µM) on steady state sarcomere length (panel A) and

2+ Ca ]i (panel B) in an individual mouse ventricular myocyte obtained from TRPV1 null mice (TRPV1-/-). Representative traces demonstrating the effect AITC (100

2+ µM) on sarcomere length (panel C) and Ca ]i (panel D) in an individual CM

obtained from a TRPV1-/- mouse. Changes in sarcomere length were measured

2+ in micrometers. [Ca ]i was measured as the 340/380 ratio.

2+ Treatment with HC030031 or SB366791 Does Not Alter [Ca ]i Dynamics or

Contractile Function in CMs

Summarized data depicting the effects of HC030031 (10 µM) and

SB366791 (10 µM) are shown in Figure 18A-F. In CMs obtained from wild-type mouse hearts, no detectable differences were observed in fractional shortening,

2+ 2+ maximum velocity of shortening/relengthening, peak [Ca ]I, time to peak [Ca ]i

or Ca2+ decay when treated with HC030031. Similarly, application of SB366791

2+ to CMs elicited no detectable differences in the [Ca ]I or contractile function

parameters listed above. Results are expressed as a percent of each treatment’s

respective control value set at 100%.

2+ Figure 18: Treatment with HC030031 or SB366791 Does Not Alter [Ca ]i

Dynamics or Contractile Function in CMs. Summarized data demonstrating the lack of an effect of HC030031 (10 µM) or SB366791 (10 µM) on contractile

2+ function and [Ca ]i cycling in CMs obtained from wild-type mouse hearts.

Results are expressed as percent of steady state baseline control (Ctrl) value set

2+ at 100%. Changes in peak [Ca ]i are measured as the change in the 340/380 ratio. Changes in timing are measured in milliseconds. * P < 0.05 compared with

Ctrl. n = 6 cells from 3 hearts.

2+ AITC- and Capsaicin-Induced Increases in CM [Ca ]i and Contractile

Function are Similar to Those Observed Following β-Adrenergic Receptor

(β-AR) Stimulation with ISO

For comparison, we also assessed the dose-dependent effects of β-AR

2+ stimulation with isoproterenol (ISO; 1-100 nM) on [Ca ]i and contractile function

in CMs. The dose-dependent effects of AITC (1-300 µM) and capsaicin (0.1-1

2+ µM) on CM [Ca ]i and contractile function were qualitatively and quantitatively very similar to those observed with ISO (1-100 nM). The summarized data are

depicted in Table 1.

AITC (n=18) Control 1 µM 10 µM 100 µM 300 µM Fractional Shortening 4.5 ± 0.1 5.0 ± 0.2 6.3 ± 0.2* 8.7 ± 0.3* 10.8 ±0.3* (% sarcomere length) Maximum Velocity of 4.7 ± 0.1 5.4 ± 0.2 7.1± 0.3* 9.9 ± 0.3* 11.2 ± 0.3* Cell Shortening (µm/sec) Maximum Velocity of 3.6 ± 0.1 4.1 ± 0.1 5.1 ± 0.3* 7.1 ± 0.3* 8.3 ± 0.3* Cell Relaxation (µm/sec) 2+ Peak [Ca ]i 0.13 ± 0.02 0.15 ± 0.04 0.19 ± 0.04* 0.22 ± 0.06* 0.26 ± 0.04* (change in 340/380 ratio) 2+ Tp [Ca ]I (msec) 158 ± 9.4 144 ± 8.4 112 ± 9.0* 58 ± 6.1* 46 ± 3.3 2+ [Ca ]i Decay (msec) 610 ± 19 563 ± 18 458 ± 15* 326 ± 12* 235 ± 9*

Capsaicin (n=12) Control 0.1 µM 1 µM

Fractional Shortening 4.7 ± 0.3 8.7 ± 0.4* 10.7 ± 0.6* (% sarcomere length) Maximum Velocity of 4.2 ± 0.2* 7.5 ± 0.5* 8.9 ± 0.5* Cell Shortening (µm/sec) Maximum Velocity of 3.4 ± 0.3* 6.7 ± 0.3* 8.4 ± 0.1* Cell Relaxation (µm/sec) 2+ Peak [Ca ]i 0.10 ± 0.03 0.20 ± 0.04* 0.21 ± 0.07* (change in 340/380 ratio) 2+ Tp [Ca ]I (msec) 147 ± 10.3 69 ± 4.4* 54 ± 4.2*

2+ [Ca ]i Decay (msec) 620 ± 19 353 ± 6.2* 198 ± 11*

ISO (n=15) Control 1 nM 5 nM 10 nM 100 nM

Fractional Shortening 4.6 ± 0.2 5.5 ± 0.3 7.1 ± 0.3* 10.7 ± 0.6* 11.5 ±0.5* (% sarcomere length) Maximum Velocity of 4.7 ± 0.1 5.1 ± 0.1 7.4 ± 0.2* 11.1 ± 0.4* 12.7 ± 0.4* Cell Shortening (µm/sec) Maximum Velocity of 3.6 ± 0.1 4.0 ± 0.1 5.6 ± 0.2* 8.6 ± 0.3* 10.0 ± 0.2* Cell Relaxation (µm/sec) 2+ Peak [Ca ]i 0.11 ± 0.02 0.12 ± 0.03 0.17 ± 0.05* 0.22 ± 0.05* 0.23 ± 0.05* (change in 340/380 ratio) 2+ Tp [Ca ]I (msec) 140 ± 17 124 ± 16 104 ± 9 56 ± 7* 52 ± 6* 2+ [Ca ]i Decay (msec) 615 ± 16 586 ± 15 381 ± 16* 231 ± 21* 174 ± 16*

Table 1: Comparison of AITC-, capsaicin- and ISO-induced changes in CM

2+ [Ca ]i and contractile function. Data are expressed as mean ± SEM. * = P <

0.05.

TRPA1 activation with AITC dose-dependently increases ejection fraction in mouse hearts

Figure 19 demonstrates the dose-dependent effects of TRPA1 stimulation

with AITC on ejection fraction in a mouse heart in vivo. Echocardiographic

assessment of left ventricular diastolic and systolic function demonstrated that

AITC (1-200 µg/kg/min) dose-dependently increased ejection fraction in wild-type

mouse hearts from 42 ± 3.4% to 73 ± 2.1% without significantly raising heart rate

(data not shown).

Figure 19: TRPA1 activation with AITC dose-dependently increases ejection fraction in wild-type murine hearts. P < 0.05 compared to baseline control value.

DISCUSSION

Our findings from Chapter two demonstrate the functional expression of

TRPA1 and TRPV1 in the adult mouse heart and their co-localization in

costameres, z-discs and intercalated discs of mouse CM’s. To our knowledge,

the current data in Chapter three are the first to thoroughly characterize the

2+ effects of TRPA1 and TRPV1 stimulation on [Ca ]i and contractile function in

electrically-stimulated adult mouse CMs. The major finding of the current study

is that TRPA1 agonist, AITC, and TRPV1 agonist, capsaicin, stimulate dose-

2+ dependent increases in peak [Ca ]i and contractile function in freshly isolated

CM’s. Moreover, a dose-dependent acceleration in the timing parameters associated with the rise and fall of the Ca2+ transient as well as shortening and

relengthening of the CM were also observed. The effects of AITC were not

observed in the presence of the TRPA1 antagonist, HC-030031 nor in CMs

-/- 2+ obtained from TRPA1 mice indicating the effects of AITC on [Ca ]i and contractile function were TRPA1 dependent. Furthermore, the capsaicin-induced

2+ increases in [Ca ]i and contractile function were absent in the presence of the

TRPV1 antagonist, SB366791, as well as in CMs obtained from TRPV1-/- mice indicating the effects of capsaicin were dependent upon TRPV1. The effects of

2+ TRPA1 and TRPV1 stimulation on [Ca ]i and contractile function were qualitatively and quantitatively similar to the classical CM responses observed

97 following β-AR stimulation with isoproterenol. Finally, TRPA1 stimulation with

AITC dose-dependently increases ejection fraction in murine hearts.

AITC and Capsaicin Stimulate TRPA1- and TRPV1-Dependent Increases in

2+ [Ca ]i and Contractile Function in CMs, Respectively

2+ Modulation of CM [Ca ]i throughout the excitation contraction coupling process, as well as the timing parameters associated with intracellular Ca2+ handling is a critical determinant of the inotropic and lusitropic state of the heart under physiological and/or pathophysiological conditions. The discovery of novel ligand gated ion channels capable of activating signaling pathways which modulate Ca2+ regulatory pathways and/or myofilament Ca2+ sensitivity may pave the way for the design of novel therapies capable of increasing the inotropic and or lusitropic state of the heart. Our findings indicate that TRPA1 and TRPV1 stimulation results in the activation of a signaling pathway(s) that modulate cellular mechanisms leading to (1) an increase in the amount of cytosolic Ca2+ available to interact with the cardiac myofilaments; (2) an acceleration in the time

2+ 2+ to peak [Ca ]i and (3) an acceleration in the removal of Ca from the cytosol.

These changes in intracellular Ca2+ availability and handling are also reflected as changes in CM contractile function including (1) an increase in fractional shortening; (2) an increase in maximum velocity of shortening and (3) an increase in maximum velocity of relengthening.

Potential Cellular Signaling Pathway(s) and Cellular Mechanisms of TRPA1-

2+ and TRPV1-Induced Increases in CM [Ca ]i and Contractile Function

Based on the current findings, we hypothesize that stimulation of TRPA1

or TRPV1 ion channels may be coupled to the protein kinase A (PKA) signaling

pathway in CMs. Our hypothesis is based on the fact that the current study

demonstrates remarkable qualitative and quantitative similarities when

2+ comparing the effects of TRPA1 or TRPV1 stimulation on CM [Ca ]i and contractile function to those effects observed in response to β-AR stimulation

(Bers, 2001; Solaro, 2011). The β-AR signaling pathway has been well established in heart for decades and is known to be coupled to the cAMP/PKA signaling pathway resulting in phosphorylation and activation of sarcolemmal

Ca2+ channels (Catterall, 1988; Hess et al., 1986) and the sarcoplasmic

reticulum (SR) pump regulatory protein, phospholamban (Katz et al., 1975; Li et

al., 2000; Tada and Inui, 1983). Moreover, it is also well established that β-AR

stimulation of intact hearts or isolated CMs results in phosphorylation of troponin

I (TnI) on the cardiac myofilaments (Kranias et al., 1985; Li et al., 2000; Onorato

and Rudolph, 1981). We propose that TRPA1 and TRPV1 stimulation likely

involves an intracellular signaling pathway(s) that trigger post-translational

modifications (phosphorylation) of the L-type Ca2+ channel as well as

phospholamban resulting increases in transsarcolemmal Ca2+ influx, SR uptake

and Ca2+ loading. Together these would, at least in part, account for an increase

in the amount of Ca2+ available to interact with the cardiac myofilaments and the

accelerated rate of decay of the Ca2+ transient due to the removal of inhibition of

the SR Ca2+ pump by phosphorylated phospholamban. Parallel changes in contractility could also be explained by the aforementioned mechanisms and include an increase in (1) fractional shortening, (2) maximum velocity of shortening and (3) maximum velocity of relaxation. Alternatively, there could also be a role for TRPA1- or TRPV1-dependent phosphorylation of TnI in mediating the observed changes in both contraction and relaxation that are independent of changes in Ca2+ handling (Solaro, 2011). It is well established that PKA-

dependent phosphorylation increases cross bridge cycling rate and maximum

unloaded shortening velocity (Vm) which contributes to the lusitropic effects of β-

AR stimulation (Hoh et al., 1988; Strang et al., 1994). We observed TRPA1- and

TRPV1-induced increases in maximum unloaded shortening velocity which could

be explained by TnI phosphorylation at PKA-dependent sites. Moreover, an

increased shortening velocity could contribute to the increased fractional

shortening and a positive inotropic effect since, in theory, the power output of

muscle is determined by the product force of velocity (Herron et al., 2001;

Layland et al., 2004). Finally, TnI phosphorylation at PKA-dependent sites is also

known to reduce myofilament Ca2+ sensitivity and contribute to an accelerated

rate of relaxation also contributing to a positive lusitropic effect (Solaro, 2011).

In vivo Effects of TRPA1 Stimulation via Acute AITC Infusion on Cardiac

Function

TRPA1 has recently emerged as a target of interest in the regulation of

cardiovascular physiology and pathophysiology (Inoue et al., 2006; Inoue et al.,

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2009; Yue et al., 2015). However, research involving TRPA1 in cardiac muscle tissue has not progressed due to the previously unknown presence of the ion channel in CMs. We reported in Chapter two the precise functional expression of

TRPA1 ion channels in mouse ventricular tissue. The next logical step was to determine the extent to which TRPA1 regulates and/or modulates contractility. In

Chapter three, we determined that TRPA1 stimulation elicits positive inotropic and lusitropic effects in individual cardiomyocytes whereas TRPA1 blockade had no significant effects. The increase in contractile function observed in vitro prompted the hypothesis that TRPA1 stimulation in vivo would result in a similar positive inotropic and lusitropic outcome. The current results demonstrate the remarkable dose-dependent increases in ejection fraction observed following

AITC infusion in wild-type mice. Furthermore, the effects on heart rate and mean arterial pressure were negligible. The novel phenomenon by which AITC induces robust increases in ejection fraction may have significant therapeutic implications whereby TRPA1 stimulation may be used to improve diminished cardiac function, such as that observed in heart failure.

Summary and Conclusions of Chapter 3

Our findings from Chapter three are among the first to identify TRP channels (TRPA1 and TRPV1, specifically) as potential targets to modulate beat- to-beat physiology of CMs. Our key findings are that TRPA1 and TRPV1 stimulation induces a robust positive inotropic and lusitropic effect in electrically- paced ventricular cardiomyocytes. Furthermore, the mechanisms by which AITC

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and capsaicin act are independent of TRPV1 and TRPA1, respectively. These

results suggest that TRPA1 or TRPV1 ion channels may serve as viable targets

to increase cardiac output in vivo. Historically, modulation of the mechanisms

2+ 2+ that regulate [Ca ]I handling and myofilament Ca sensitivity are primarily

attributed to SNS stimulation and the resultant release of catecholamines. In fact,

catecholamine-induced β-adrenergic receptor (β-AR) activation is widely

regarded as the most powerful physiological mechanism to increase cardiac

performance. Stimulation of the β-ARs results in activation of the Gs signal transduction pathway, which includes adenylyl cyclase, cAMP and PKA. PKA subsequently phosphorylates a myriad of downstream effectors, the main targets of which are the following: 1) the sarcolemmal LTCC and SR RYR receptors,

2+ each of which will induce increases in [Ca ]I; 2) PLB, whose phosphorylation

accelerates Ca2+-reuptake through the SERCA into the SR; 3) TnI and myosin

binding protein-C, each of which modulate the relaxation of cardiac muscle and

myofilament Ca2+ sensitivity and 4) phospholemman, a Na+/K+-ATPase inhibitor,

whose phosphorylation removes the inhibition and stimulates the Na/K pump,

effectively promoting the rate of cardiac muscle repolarization and subsequent

relaxation.

Taken together, these PKA-mediated phosphorylation events regulate CM

Ca2+-handling and, through myofilament Ca2+ sensitivity regulation, are able to

modulate inotropic (force of myocardial contraction), chronotropic (rate of firing

through the sinoatrial node/heart rate) and lusitropic (rate of myocardial

relaxation) effects on contractility. Determining the extent to which TRPA1 or

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TRPV1 stimulation modifies these downstream effectors will be the focus of future investigations. In fact, the current findings mirror those observed with β-AR

stimulation of the cAMP/PKA signaling pathway and suggest a similar pathway

2+ may be involved in TRPA1- or TRPV1-dependent changes in [Ca ]i and

contractile function of CMs. Additionally, TRPA1 and TRPV1 blockade did not

alter intracellular calcium concentrations or contractile function in electrically-

stimulated CMs. This suggests that TRPA1 and TRPV1 ion channels do not

regulate beat-to-beat contraction within cardiomyocytes.

Finally, we demonstrated that TRPA1 stimulation in vivo induces a dose- dependent increase in ejection fraction in wild-type mice while having negligible effects on heart rate and mean arterial pressure; this suggests that TRPA1 may serve as a viable target to increase ejection fraction in certain pathological conditions where cardiac output is compromised, such as congestive heart failure. In conclusion, the current results are the first to report a potential role of

TRPA1 and TRPV1 in modulating CM physiology by which their stimulation elicits robust increases in fractional shortening and peak calcium amplitude, as well as accelerating the maximal velocity of shortening and relengthening, time to peak calcium and the calcium decay rate.

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CHAPTER FOUR

The role of TRPA1 in myocardial infarction (MI) and ischemia-induced cell

death

INTRODUCTION

Compelling epidemiological and clinical data indicate that heart disease is the leading cause of death in the United States, claiming more lives than all forms of cancer combined. In the United States, someone has a heart attack every 34 seconds. Annually, direct and indirect costs of heart disease-related health expenses total more than $30 billion. Despite the intense number of studies already performed, the molecular basis of heart disease remains elusive.

The need for new and more effective therapeutic strategies to decrease morbidity and mortality are essential for improved patient outcomes and to reduce health- related expenses in the future.

TRPA1 and TRPV1 channels are members of the TRP channel superfamily of structurally related, non-selective cation channels that tend to show high permeability to calcium (Fernandes et al., 2011). Previous studies

104

have demonstrated that these channels communicate with each other to a

considerable extent in sensory neurons and other heterologous expression

systems (Patil et al., 2010; Ruparel et al., 2008; Story et al., 2003), where they function as sensory transducers of noxious stimuli (Caterina et al., 1997; Palazzo

et al., 2008). Investigations conducted by our laboratory have demonstrated that

TRPA1 agonists restore previously desensitized TRPV1 receptor sensitivity via

PKCε-dependent phosphorylation of TRPV1 in mouse sensory neurons (Wickley et al., 2010). Furthermore, we recently outlined the mechanism by which TRPA1 activation stimulates PKCε activation/phosphorylation and subsequent TRPV1 phosphorylation (Zhang et al., 2011) - an effect that was shown to occur via a nitric oxide synthase-dependent pathway (Sinharoy et al., 2015). Indeed, TRPA1 and TRPV1 are extensively co-expressed in different tissue types throughout the body, including the cardiovascular system (Kaneko and Szallasi, 2013;

Watanabe et al., 2008), where they have been implicated in vasoconstriction, vasodilation (Inoue et al., 2006) and numerous cardiovascular disorders (Inoue et al., 2009; Pozsgai et al., 2010). Recent evidence suggests the potential role of various members of the TRP superfamily to be involved in regulating overall cardiac function and cardioprotection following ischemia (Guinamard et al., 2014;

Maa et al., 2015; Xie et al., 2012). Although the specific roles of TRP channels on cardioprotection have yet to be fully elucidated, the presence of TRPV1 in cardiac tissue has been shown to provide cardioprotection following ischemia

(Huang et al., 2009). Additionally, we identified the presence of TRPA1 channels in CMs in Chapter two of this dissertation; however, the extent to which TRPA1 is

105

involved in pathophysiological remodeling processes, such as those observed

after myocardial infarction, have yet to be determined.

In the current studies, we tested the hypothesis that TRPA1 gene deletion

would result in exacerbated LV remodeling and worsened cardiac function

following MI. Furthermore, we hypothesized that TRPA1 stimulation with AITC

would induce a cardioprotective mechanism involving eNOS, PKCε and TRPV1.

MATERIALS AND METHODS

Animal Models

4-month-old male WT and TRPA1-/- mice (n=4/group) were utilized and

maintained in accordance with the Guide for the Care and Use of Laboratory

Animals (NIH). Kent State University animal care facility (Kent, OH) housed all animals, which is accredited by the American Association for Accreditation of

Laboratory Animal Care.

Induction of Myocardial Infarction

MI procedures were performed as previously described (Luther et al.,

2013). In brief, four-month-old WT and TRPA1-/- male mice (Jackson

Laboratories, Bar Harbor, ME) were subjected to left anterior descending (LAD)

artery permanent occlusion or a sham surgery. Mice were injected with atropine

sulfate (0.04 mg/kg, i.m.) and anesthetized with ketamine/xylazine in saline (100

106

mg/kg) before being moved to a small rodent SurgiSuite (Kent Scientific

Corporation) and monitored via ECG probes while body temperature was

maintained at 37°C utilizing a rectal probe. Mice were tracheally intubated with a

20g fiber optic angiocatheter and connected to a small rodent ventilator (Minivent

type 845; Harvard Apparatus). A left thoracotomy (through the fourth and fifth

intercostal space) was carried out and occlusion of the LAD artery was

performed using an 8-0 nylon suture. MI was confirmed by ST segment elevation

and apex blanching. The thoracic cavity was closed with a 5-0 Vicryl suture.

Animals were moved to an isolated heated recovery area and supplemented with

100% oxygen. Sham-operated mice underwent the same process except for LAD

artery ligation.

Risk Area, Fibrosis and Infarct Size (Immunohistochemical Studies)

Histological assessment of Masson’s trichrome was carried out as

described previously (Luther et al., 2012). Infarct size was analyzed utilizing

2,3,5-triphenyltetrazolium chloride (TTC) following MI. Hearts were subsequently excised and not sectioned (whole heart) or sectioned at the mid-ventricular and

apex levels (2 mm transverse slices). Sections were incubated in TTC for 20

minutes at 37°C (viable = red and necrotic = white) and incubated in 4% PFA for

20 minutes at room temperature. To determine density of collagen deposition,

Masson’s trichrome-stained cells were counted in 5 different fields of the peri-

infarct zone under a microscope and reported as a percent of collagen in the

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ventricle. Heart sections were assessed using ImageJ Software (NIH, Bethesda,

MD).

Transthoracic Echocardiography

In vivo cardiac functional parameters were assessed utilizing a Vevo 770

(VisualSonics, Inc., Toronto, Ontario, Canada) in the following animal groups:

wild-type mice before and seven days following myocardial infarction. Mice were anesthetized using 2% isoflurane and placed on an adjustable platform equipped with ECG electrodes to monitor heart and respiration rates. Doppler echocardiography was carried out parasternally across a shaved chest wall and short (midpapillary level)- and long-axis images were obtained. Functional measurements were averaged from three cardiac cycles. The following parameters were assessed as indicators of cardiac function: LV diastolic internal diameter (LVIDd), LV systolic internal diameter (LVIDs), posterior wall thickness and ejection fraction (LVIDd-LVIDs/LVIDs × 100). Measurements were calculated blinded reviewers using the Vevo 770/3.0 software.

CM Isolation

Murine hearts were excised, prepared for aortic cannulation and transferred to a Langendorff apparatus for CM isolation, as previously described

(O'Connell et al., 2007). In brief, hearts underwent retrograde perfusion at 37°C and pH 7.4 with a modified Krebs-Henseleit buffer (in mM: 120.4 NaCl, 4.8 KCl,

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0.6 KH2PO4, 0.6 Na2HPO4, 1.2 MgSO4-7HsO, 10 Na-HEPES, 4.6 NaHCO3, 30

taurine, 10 BDM, and 5.5 glucose). The Krebs-Henseleit buffer was sterile- filtered and paced at a rate of 4 mL/min. After perfusion for 4 minutes, the digestion buffer containing collagenase type II (300 U/mg, Worthington

Biochemical, Lakewood, NJ) perfused the heart for an additional 8 minutes until the heart became ‘spongy’. The left ventricles were removed, minced, and triturated in Krebs-Henseleit buffer containing fetal bovine serum. The resulting cellular digest was washed and resuspended at 23°C in HEPES-buffered saline

(in mM: 118 NaCl, 4.8 KCl, 0.6 KH2PO4, 4.6 NaHCO3, 0.6 NaH2PO4, 5.5 glucose,

pH 7.4). CM yield was ~80-90%. CMs were then untreated or treated with AITC

(100 μM). All samples were then either subjected to immunoblotting or prepared for in vitro ischemia preparations.

Preparation of Cell Lysates for Immunoblot Analysis

Immunoblot analysis was performed as previously described (Sinharoy et

al., 2015). CMs were homogenized and protein concentration was subsequently

assessed using the Bradford method (Bradford, 1976). All samples were

adjusted to ~2 mg/mL protein concentration. Samples containing 50 µg of protein

lysates were boiled and subjected to SDS-PAGE on 4-15% precast

polyacrylamide gels (Bio-Rad) through the use of a minigel apparatus. After

running, gels were then transferred to nitrocellulose membranes. Nonspecific

binding was blocked with 1% BSA solution in Tris-buffered saline solution (0.1%

[vol/vol] Tween-20 in 20 mM Tris base, 137 mM NaCl, pH 7.6) for 1 hour at room

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temperature. Antibodies recognizing pro-caspase and cleaved caspase 3

(ThermoFisher) and total GAPDH (Millipore) were diluted 1:1000 in Tris-buffered saline containing 1% BSA and incubated at 4°C overnight. After washing, membranes were incubated for 1 h at room temperature with horseradish- peroxidase linked secondary antibody (goat anti-rabbit and goat anti-mouse) diluted 1:5000 in Tris-buffered saline with 1% BSA. Enhanced chemiluminescence was used for antibody detection utilizing an ImageQuant

LAS 4000 Mini (General Electric). Immunoreactivity was assessed by scanning densitometry and analyzed using ImageJ software (NIH).

In vitro ischemia-mimetic protocol

Freshly isolated CMs obtained from wild-type and TRPA1-/- mice were

incubated in a buffer designed to simulate the ischemic environment during MI

(Esumi et al., 1991; Zhao et al., 2014).

Statistical Analysis

All experimental protocols were repeated in a minimum of four different

mice. Within group comparisons were made using one-way ANOVA and

Bonferroni post hoc test. Differences were considered statistically significant at p

< 0.05. All results are expressed as mean + SEM. Statistical analysis was

conducted using Sigmaplot 11.0 (Systat software).

110

RESULTS

TRPA1-/- mice demonstrate exacerbated scar formation and collagen deposition compared to wild-type mice following myocardial infarction

Figure 20 demonstrates the extent to which TRPA1-/- mice undergo

aberrant scar formation following myocardial infarction. Hearts obtained from

TRPA1-/- mice following LAD ligation for seven days displayed worsened left

ventricular anterior wall integrity (Figure 20A) and scar formation at the mid- ventricular level (Figure 20B) as well as at the apex level (Figure 20C) when compared to wild-type mouse hearts. Infarct/LV area ratio was significantly elevated in TRPA1-/- MI mice compared to wild-type (22 ± 7.0% vs 16 ± 3.0%;

Figure 20D). Quantitative analysis of collagen content in the LV peri-infarct zone was visualized using Masson’s trichrome staining (Figure 20E). Total collagen volume (percent collagen/LV area) was significantly exacerbated in TRPA1-/-

mice compared to wild-type (22 ± 5.0% versus 11 ± 5.1%; Figure 20F).

Differences in total collagen volume of wild-type and TRPA1-/- sham mice were

not statistically significant (data not shown).

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Figure 20: TRPA1 gene deletion leads to exaggerated scar formation following myocardial infarction in mice. Representative whole heart images demonstrating

112

LV wall integrity is compromised in TRPA1-/- mice compared to WT mice 7d

following MI (A). Representative heart sections at the mid-ventricle (B) and apex levels (C) stained with 2,3,5-triphenyltetrazolium (TTC) to show exaggerated infarct/LV area ratio in TRPA1-/- mice. Summarized data comparing infarct/LV area ratios at the apex (D). Histological assessment via Masson’s trichrome staining (400x; (E)) shows increased collagen deposition (blue staining) in

TRPA1-/- mice. Summarized data depicting collagen volume in non-MI, WT 7d following MI and TRPA1-/- 7d following MI (F). * P < 0.05 compared to wild-type

mean control value. n = data obtained from 4 hearts.

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Cardiac function is deteriorated in TRPA1-/- mice compared to wild-type following MI

Figure 21 depicts the deteriorated cardiac function in TRPA1-/- mice

compared to wild-type before and seven days after MI. Representative left

ventricular echocardiography recordings demonstrate exaggerated anterior wall

dyskinesis seven days after MI in TRPA1-/- mice when compared to wild-type.

Baseline recordings and sham-operated (data not shown) TRPA1-/- and wild-type

mice were not significantly different. Quantitative analysis of 2-dimensional

guided M-mode tracings show deteriorated chamber dimension, fractional area

change and ejection fraction in TRPA1-/- MI mice (Table 2), illustrating the

augmented cardiac function in wild-type MI mouse hearts.

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WT WT 7d MI TRPA1-/- TRPA1-/- 7d MI

BW (g) 42.3±3.1 37.8±3.2 41.9±3.0 37.3±4.0

HR (bpm) 504±3 549±12* 507±5 558±17**

LVIDd (mm) 4.3±0.4 4.7±0.6* 4.2±0.2 4.9±1.1**

LVIDs (mm) 2.9±0.1 3.7±0.4* 2.8±0.1 4.0±0.5**

FAC (%) 46.1±3.8 32.1±2.7* 45.8±4.1 27.3±4.5**

EF (%) 72.3±2.2 45.6±5.7* 71.9±3.1 39.3±4.3**

Figure 21 and Table 2: TRPA1-/- mice exhibit deteriorated cardiac function following MI. Representative parasternal short axis M-mode tracings of WT and

TRPA1-/- mouse hearts taken at the LV mid-papillary level during diastole and systole to show greater wall kinesis before and 7 days (7d) following MI.

Summarized echocardiography data from WT, TRPA1-/- and TRPAV-/- before and 7d following MI (Table 2). Body weight (BW), heart rate (HR), left ventricular internal diameter, diastole (LVIDd), left ventricular internal diameter, systole

(LVIDs), fractional area change (FAC), and left ventricular ejection fraction (EF). *

P < 0.05 compared to WT before induction of MI. ** P < 0.05 compared to wild- type 7d following MI. n = data obtained from 4 hearts.

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TRPA1 stimulation attenuates ischemia-induced cell death through a signal

transduction mechanism dependent on eNOS and PKCε, but not TRPV1, in

CMs

Figure 22 demonstrates the effects of TRPA1 stimulation on ischemia-

induced cell death in CMs obtained from wild-type, TRPA1-/-, NOS-/-, PKCε-/- and

TRPV1-/- mice over the course of three hours. Initially, a control experiment was

conducted with wild-type CMs exposed to the ischemia-mimetic buffer to outline

the time frame of cell death in our in vitro preparation (Figure 22A). TRPA1

stimulation with AITC attenuated ischemia-induced cell death in CMs obtained

from wild-type (Figure 22B-E) and TRPV1-/- (Figure 22E) mice in as early as the

first hour, but not TRPA1-/- (Figure 22B) NOS-/- (Figure 22C) or PKCε-/- (Figure

22D) mouse hearts. Furthermore, the rate at which TRPA1-/-, PKCε-/- and TRPV1-

/- CMs died was accelerated compared to wild-type under untreated and treated

conditions in as early as the first hour. This effect was absent in CMs obtained

from NOS-/- mouse hearts. Data were calculated as the ratio of pro-caspase 3

(pre-apoptotic marker) to cleaved caspase (post-apoptotic marker) and are expressed as a percent of the 0 hour control.

116

Figure 22: AITC attenuates ischemia-induced CM cell death. Analysis of ischemia-induced progression to CM death over the course of three hours

117

demonstrates that AITC acts through a TRPA1/eNOS/PKCε-dependent pathway.

Representative immunoblots depicting the rate at which wild-type CMs undergo

ischemia-induced cell death in the untreated condition (A). Further analysis

demonstrating ischemia-induced cell death in CMs obtained from wild-type (B-E),

-/- -/- -/- -/- TRPA1 (B), NOS (C), PKCε (D) and TRPV1 (E) mouse hearts in the

presence and absence of TRPA1 agonist, AITC (100 µM). Data are expressed as

a percent of pro-caspase 3 (pre-apoptotic marker) to cleaved caspase (post- apoptotic marker) at 0 hours. GAPDH was probed as the loading control. * P <

0.05 compared to untreated wild-type CMs at corresponding time point. ƚ P < 0.05

compared to wild-type CMs treated with AITC at corresponding time point. n =

data obtained from 4 hearts.

118

DISCUSSION

To our knowledge, the current data in Chapter four is the first to demonstrate that TRPA1 deficiency results in exaggerated cardiac fibrosis, deteriorated cardiac function and increased cell death following ischemia. The cellular and molecular events underlying aberrant cardiac remodeling and diminished function may involve excessive collagen deposition and diminished levels of crucial cardioprotective intracellular mediators in TRPA1-/- mice. Our key

findings are: 1) TRPA1 gene deletion exacerbates scar formation and atypical

cardiac remodeling following myocardial infarction, 2) TRPA1 gene deletion leads

to compromised myocardial function following ischemia and 3) TRPA1-induced

attenuation of ischemia-induced CM death involves an eNOS/PKCε-dependent

pathway.

Excessive Scar Formation and Diminished Cardiac Function in TRPA1-/-

Mice

Two crucial prognosis factors commonly used to assess the severity of

myocardial injury following MI deal with the size of infarction and LV remodeling.

We found that TRPA1-/- mice exhibit excessive scar formation accompanied by

elevated collagen deposition and compromised left ventricular anterior wall

integrity. This suggests that TRPA1 may be involved in collagen degradation

119

and/or deposition which may lead to ventricular rupture in severe infarctions. The

severity of MI and pathological collagen deposition dictates the extent to which

cardiac function is compromised. Increased collagen deposition alters ventricular

compliance, inducing increases in LV stiffness and diminished LV performance

that lead to the progression toward congestive heart failure and mortality (Huang et al., 2009). Indeed, we found that exaggerated scar formation in TRPA1-/- MI mice reduced the normal functioning of the myocardium, rendering the resultant cardiac functional parameters deteriorated when compared to wild-type MI mice.

The most significant alteration dealt with the altered ejection fraction which was reduced nearly ~30% in TRPA1-/- MI mice whereas wild-type experienced only a

~20% decrease.

Notably, TRPA1 has recently been identified in cardiac fibroblasts

(Pazienza et al., 2014); therefore, alterations in LV remodeling and fibrosis patterns in TRPA1-/- MI mice cannot be solely attributed to the lack of TRPA1 ion

channels in CMs. In fact, the absence of TRPA1 in fibroblasts may affect the

efficiency by which products of oxidative stress induce fibroblast proliferation

and/or differentiation to myofibroblasts rendering the heart subject to increased

risk of aberrant remodeling. As such, data obtained from global knockout mouse

in in vivo investigations should be interpreted with caution due to the high

prevalence of systemic compensatory mechanisms taking place. To confirm the trends observed in TRPA1-/- MI mice, further investigations should be carried out utilizing TRPA1 antagonists or TRPA1 knock-down models.

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Delineating a TRPA1-Mediated Mechanism of Cardioprotection

Previous studies from our lab and others have shown that TRPA1 and

TRPV1 may cross-talk/interact through distinct yet complex intracellular signaling

cascades that involve an eNOS/NO/PKCε-dependent signal transduction

mechanism (Sinharoy et al., 2015). As such, we hypothesized that TRPA1

stimulation would result in a similar mechanistic outcome in CMs. We

subsequently employed an in vitro approach to determine the extent to which the

TRPA1/eNOS/PKCε/TRPV1 pathway modulates ischemia-induced cell death.

We created an ischemia-mimetic buffer designed to simulate the extracellular

environment that would be observed in the in vivo setting and incubated primary

isolated CMs obtained from wild-type, TRPA1-/-, NOS-/-, PKCε-/- and TRPV1-/-

mice in the presence or absence of AITC. The current results suggest that

TRPA1 stimulation attenuates ischemia-induced cell death in wild-type and

TRPV1-/- CMs but not in those obtained from TRPA1-/-, NOS-/- or PKCε-/-.

Previous studies in the heart have demonstrated that activation of PKC

reduces MI injury, whereas PKC inhibition abrogates ischemic preconditioning

(Cohen and Downey, 1995; Ytrehus et al., 1994). Moreover, it has also been

shown that this cardioprotective effect can be fully mimicked by modulating the

activity of the ε isoform, however evidence also points towards the delta (δ)

isoform of PKC (Gray et al., 1997; Hao et al., 2014; Liu et al., 1999). Multiple

molecular events have been shown to have an activating effect on PKCε, the

most important of which is NO. Although the effects of NO on PKCε depend on

its biological functions and cell type (Nishio and Watanabe, 1997; Tepperman et

121 al., 2009; Yoshida et al., 1999), NO-induced PKC activation in cardiac tissue is well established (Liu et al., 2001). Exogenous NO (released by NO donors) has been shown to induce PKCε activation (Bolli, 2001; Cohen et al., 2000; Xi and

Kukreja, 2000). Furthermore, activation of PKCε has been demonstrated to play an essential role in mediating signal transduction events during NO-induced cardioprotection following ischemia (Balafanova et al., 2002). Moreover, eNOS has been implicated in serving a variety of roles within cardiac tissue including myocardial protection following ischemic insult (Kukreja and Xi, 2007; Scherrer-

Crosbie et al., 2001), regulation of caveolin-3-mediated protection from hypertrophic cardiomyopathy (Park et al., 2002; Wooman et al., 2002), the production of superoxide anions (Pritchard Jr et al., 2001) and potentially mediating the cardiac inotropic response to sustained stretch (Eisner et al.,

2000). The polymodal activation mechanisms and intricate regulatory functions of eNOS and PKCε suggest a notable complexity of the cellular signaling pathways in which they are involved. As such, elucidating signal transduction mechanisms activating eNOS and PKCε in cardiac tissue are of the utmost importance and require further experimentation.

It should be noted that although evidence exists describing the cellular signal transduction pathways and molecular mechanisms in which multisite eNOS phosphorylation occurs in cardiac tissue (Fleming, 2010; Kukreja and Xi,

2007; Massion et al., 2003; Mount et al., 2007), the current understanding of the physiological and pathophysiological implications remain uncertain. However, eNOS phosphorylation at serine 1177 is widely accepted as an indicator of

122 enzyme activity. In order to elucidate the precise physiological and pathophysiological implications underlying alterations in phosphorylated eNOS levels in cardiac tissue, future experiments will be required; however, the current results have prompted several hypotheses. First, phosphorylated eNOS has been implicated in serving a role in cardioprotection following ischemic insult in the heart (Kukreja and Xi, 2007; Scherrer-Crosbie et al., 2001); therefore, altering expression levels of phosphorylated eNOS may confer a cardioprotective phenotype to protect against exacerbated cardiac injury and atypical remodeling following myocardial infarction or ischemia-reperfusion injury. Circumstances where diminished phosphorylated eNOS expression levels are observed may also confer a potential phenotypic alteration of NO handling in cardiac tissue.

Second, the protective regimens commonly used to minimize the irreversible injury induced by acute myocardial ischemia, such as statins, have been demonstrated to act via the pro-survival Akt-eNOS pathway (Bell and Yellon,

2003; Birnbaum et al., 2005; Wolfrum et al., 2004). Third, eNOS phosphorylation has been demonstrated to be protective against apoptosis in myocardial tissue

(Gao et al., 2002). Notably, previous studies indicate that eNOS-/- mice exhibited only a moderate attenuation of the cardioprotective effects afforded by preconditioning (Bell and Yellon, 2001). This suggests that compensatory mechanisms may be occurring in eNOS-/- mice, such as those offered by inducible (iNOS) and neuronal NOS (nNOS).

Recent studies have identified PKCε as a downstream effector of TRPA1 in tissues around the body (Sinharoy et al., 2015). To our knowledge, the current

123

data are the first to suggest a downstream effect of PKCε as a result of TRPA1

stimulation in CMs. The therapeutic implications of PKCε are vast; however, it is

widely regarded as one of the most powerful indicators of cardioprotection in

myocardial ischemia by which it preserves CM integrity and viability following

exposure to ischemia. Therefore, we hypothesized that PKCε may be involved in

the protective response mediated by TRPA1. Gene deletion of PKCε completely

abrogated the protective effects of TRPA1 stimulation in our in vitro preparations suggesting a role in mediating the apoptotic machinery within CMs. This suggests a crucial role of PKCε as an intracellular mediator in TRPA1-induced cardioprotection. These results are consistent with numerous investigations outlining the cardioprotective nature of PKCε in cardiac tissue (Dorn et al., 1999;

Gray et al., 1997; Liu et al., 1999).

Additionally, the presence of TRPV1 has also been implicated in serving a cardioprotective role following ischemic events in normal and diabetic mouse hearts (Huang et al., 2009; Ren et al., 2011). The current data suggests that the absence of TRPV1 in CMs did not significantly affect TRPA1-induced CM survival in ischemia. Therefore, we concluded that TRPA1-mediated attenuation of ischemia-induced CM death occurs independently of TRPV1 in CMs.

Although the current data identifies TRPA1-mediated signaling pathways as promising therapeutic targets for ischemic heart injury, determining the extent to which TRPA1-mediated eNOS, PKCε and/or TRPV1 phosphorylation modulates physiological and pathophysiological events in cardiac tissue remain to be fully determined and require further experimentation.

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Limitations

The discussion for Chapter four is largely speculative due to the paucity of information underlying TRPA1 signaling in cardiac tissue. Indeed, further experiments will need to be carried out to fully characterize the function of

TRPA1 and TRPV1 ion channels in cardiac muscle. The current data suggests a promising role of TRPA1 signaling in cardioprotection following ischemia in the heart; however, this study has several limitations and needs to be interpreted with caution. First, data obtained from global knockout mice in in vivo investigations (i.e. TRPA1-/- MI mice) may experience neglected systemic compensatory mechanisms. To confirm the trends observed in TRPA1-/- MI mice, further investigations should be carried out utilizing TRPA1 antagonists or

TRPA1 knock-down models. Second, cell death was measured using procaspase 3 (pre-apoptotic marker) and cleaved caspase-3 (post-apoptotic marker) which are typically associated with the indication of apoptosis although both apoptosis and necrosis play a role in the process of tissue damage subsequent to MI; however, apoptosis has been demonstrated to be the major determinant of infarct size. Distinguishing the extent to which cell death in response to ischemic insult occurs via apoptosis or necrosis may provide fundamental insight into the process of cardiac degeneration following ischemic injury. Although both necrosis and apoptosis result in the death of the CM, they differ in several cell regulatory features. There is no totally specific marker that solely detects apoptotic cells, so a combination of techniques should always be used to detect and distinguish between apoptosis and necrosis. Furthermore,

125 since CM cell death resulting from ischemic insult is commonly linked to mitochondrial-regulated pathways, measurements assessing regulatory proteins could be utilized to detect Bcl-2 (anti-apoptotic/apoptosis inhibitor), Bax (pro- apoptotic) or the apoptotic marker, soluble Fas (sFas). Although Annexin V and propidium iodide are useful in flow cytometry analysis of cell death, lactate dehydrogenase release and 7-aminoactinomycin D are both useful in the detection of apoptosis and necrosis via colorimetric absorbance assays and fluorescence microscopy, respectively. Third, other cardioprotective mediators besides eNOS and PKCε may carry out specific functions downstream of TRPA1 activation in CMs or cardiac tissue as a whole. Lastly, the current studies are preliminary in nature. Extensive histological assessment will need to be carried out in order to fully characterize post-MI remodeling in TRPA1-/- mouse hearts; this includes picrosirius red staining, Evan’s blue staining, inflammatory cell infiltration and apoptotic nuclei identification via TUNEL.

Summary and Conclusions of Chapter 4

Our key findings from Chapter four are that gene deletion of TRPA1 worsens scar formation and cardiac functional parameters compared to wild-type mice following MI. The current results are consistent with previous investigations demonstrating the cardioprotective nature of several other TRP channels

(however, other investigations have demonstrated that gene deletion of TRP channels may preserve cardiac function and attenuate aberrant LV remodeling following MI). Additionally, the current data is the first to identify a potential signal

126 transduction mechanism involved in TRPA1-mediated protection from ischemia in CMs. Stimulation of the TRPA1 attenuated ischemia-induced apoptosis in in vitro preparations – a process that is dependent upon NOS and PKCε, but independent of TRPV1 in CMs. To our knowledge, this is the first known mechanism delineated for TRPA1 stimulation in CMs. The extent to which this pathway modulates other physiological or pathophysiological events in cardiac tissue will be the focus of future investigations. The polymodal activation mechanisms and intricate regulatory functions of TRPA1, eNOS, PKCε and

TRPV1 suggest a notable complexity of cellular signal transduction pathways to which they are involved. Furthering the current understanding of the mechanisms initiated via TRPA1 stimulation in cardiac tissue could provide important fundamental insight into the development of therapeutic agents designed to combat cardiac pathophysiological conditions such as congestive heart failure and myocardial infarction.

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CHAPTER FIVE

CONCLUSIONS

The current data in this dissertation is the first to outline the general characteristics of TRPA1 and TRPV1 in cardiac tissue. In Chapter two, we identified the precise ultrastructural localization of TRPA1 and TRPV1 in cardiac tissue whereby they colocalize at the Z-discs, costameres and intercalated discs throughout the endo-, myo- and epicardial layers. The localization of TRPA1 and

TRPV1 in CMs have prompted several hypotheses. First, the Z-disc is the site of localization for many proteins, which indicate that the ion channels may share similar signaling pathways and/or are involved in direct physical interactions with other structures located therein. Secondly, CMs have stress-strain sensors embedded at several locations, including the Z-disc, costameres and intercalated discs; this suggests a potential role for the receptors in mediating mechanotransduction (Hoshijima, 2006). Lastly, the localization of the channels at the intercalated discs could be correlated with the presence of proteins which mediate calcium-dependent cell-to-cell adhesion, such as N-cadherin (Li, 2014;

Sheikh et al., 2009). We also demonstrated that AITC and capsaicin elicit dose-

128 dependent, transient increases in intracellular free calcium concentrations through TRPA1- and TRPV1-dependent processes, respectively, in CMs.

In Chapter three, we identified TRPA1 and TRPV1 as potential targets to modulate cardiac contractility by increasing inotropy and lusitropy. Furthermore, the current data in Chapter three indicates that TRPA1 and TRPV1 do not regulate beat-to-beat physiological mechanisms of contracting CMs. In vivo administration of AITC also dose-dependently increased ejection fraction in mice.

Taken together, the current data in Chapter three suggests that these ion channels may serve as novel therapeutic targets to improve the pumping capacity of the heart. The translational significance of these findings lies in the potential ability of TRP ion channels to increase the diminished cardiac function observed in certain cardiac pathologies, such as congestive heart failure.

Finally, we identify TRPA1 as a crucial cardioprotective mediator in

Chapter four. The current results are consistent with previous investigations demonstrating the cardioprotective nature of several other TRP channels.

Additionally, the current data is the first to identify a potential signal transduction mechanism involved in TRPA1-mediated protection from ischemia in CMs.

Stimulation of the TRPA1 attenuated ischemia-induced apoptosis in in vitro preparations – a process that is dependent upon NOS and PKCε, but independent of TRPV1 in CMs.

Overall, the current data in this dissertation provides a foundation of knowledge from which future investigations can conjure their hypotheses regarding TRP ion channels in cardiac tissue. Our investigations suggest that

129

TRPA1 and TRPV1 may serve as viable targets for the development of therapeutic agents designed to combat diminished cardiac function and/or ischemic injury in the heart.

130

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