Role of autophagy in cardiac fibroblast activation in ... - MSpace

Role of autophagy in cardiac fibroblast

activation in cardiac fibrosis

by

Shivika Gupta

A Thesis submitted to Faculty of Graduate Studies of

The University of Manitoba

in partial fulfilment of the requirements of the degree of

Doctor of Philosophy

Department of Physiology and Pathophysiology

University of Manitoba

Winnipeg

Copyright ©2017 by Shivika Gupta

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Abstract

Following myocardial infarction (MI), initial cardiac remodeling or wound repair/healing is

beneficial and required for continued function. During the wound healing phase, cardiac

fibroblasts activate to become contractile and hypersynthetic myofibroblasts and contribute to

cardiac fibrosis. Chronic cardiac remodeling leads to the development of overt cardiac

hypertrophy and excessive extracellular matrix (ECM) deposition with attendant loss of

myocardial performance. There is no treatment that prevents or reverses fibrosis in the post-

MI heart. Induction of autophagy is linked to stress-induced ventricular remodeling in various

cardiac diseases and has not been well studied in fibroblast activation. As the plating of

primary cardiac fibroblasts on incompressible plastic substrate leads to activation of these

cells, we used this model system in primary rat cardiac fibroblasts to examine the relationship

between autophagy and fibroblast activation. 72 hours after plating of fibroblasts, we

observed the concomitant increase in expression of myofibroblast markers and induction of

autophagy, assessed via western blotting, immunofluorescence, and transmission electron

microscopy. Treatment with inhibitors of autophagy (BafilomycinA1 (Baf-A1), 3-

methyladenine (3MA), and chloroquine (CQ) decreased myofibroblast marker expression and

reduced myofibroblast contractility (assessed by collagen gel contraction assay). CQ

treatment suppressed fibroblast activation in unpassaged P0 cardiac fibroblasts. We used the

coronary artery ligation rat model of MI to further explore the effects of CQ administration in

development of cardiac fibrosis. At 4 and 8 weeks post-MI, there was no significant change

in ventricular function (assessed via echocardiography), however in isolated cardiac tissue

treated with CQ, we observed a decrease in myofibroblast marker expression. We also show

increased autophagy in the MI group animal. We also studied tissue fibrosis by comparing

the collagen accumulation between CQ treated and nontreated animals. Although CQ

treatment was associated with a decrease in -SMA expression in the 4 week post-MI group,

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there was no difference seen on collagen level among the same study groups. These data

support a causal link between autophagy and activation of cardiac fibroblast. This study

provides a new avenue for therapeutic options to suppress the activation of cardiac fibroblasts

and thereby reduce cardiac fibrosis.

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Acknowledgements

I would like to express gratitude to Dr. Ian Dixon for his support, guidance and

encouragement during the entire course of my Ph.D. training. I am grateful for his advice,

patience, and numerous opportunities he has provided me, which has helped me to become a

better researcher and more confident person. I would also like to thank my advisory

committee members including Drs. Andrew Halayko, Jeffrey Wigle, and Thomas Netticadan

for their valuable suggestions which helped to improve my research. Their patience and

efforts to enhance my critical thinking have allowed me to become a better graduate student.

I would specially like to extend my gratitude to Dr. Saeid Ghavami who has guided me

during the entire project with his expert knowledge in autophagy. I would like to thank the

CIHR, the Institute of Cardiovascular Sciences, the Wyrzykowski family and the Deacon

Foundation for supporting me. The financial support from these agencies has been critical to

my success.

I greatly appreciate our lab manager Sunil Rattan for his hands-on assistance in all the

experiments. Sunil has offered great advice professionally and personally during my training,

and has acted as both a teacher and a friend. His positive attitude and guidance was always

welcome. I am very grateful to my lab members Krista Filomeno, Dr. Matthew Zeglinski,

Morvarid Kavosh, and Natalie Landry for their support and comraderie. Scientific

discussions with all the labmates were always helpful. I would especially like to thank Krista

Filomeno, who has been an amazing friend and confidante. I would like to extent my

gratitude to the members of Kardami lab, Navid Kholini and Robert Fandrich for use and

instruction of equipment, and always providing their expertise when needed. I feel extremely

honored to be a part of the St. Boniface Hospital Albrechtsen Research Centre, and the

Institute of Cardiovascular Sciences. I would also like to give a special thank you to Dr.

Pawan Singal for his constant support and guidance to graduate students in the Institute. I

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would also like to thank Dr. Singal’s lab members for sharing equipment, reagents, and

friendship, and to the office ICS staff (Mary Brown and Shweta Sharma), who are always

available to help with administrative issues and always have a smile on their faces. I also

extend my thanks to Drs. Peter Cattini (Head of the Department of Physiology and

Pathophysiology) and Janice Dodd.

Finally, I would like to thank my family and friends. Without their love and support I

would not have reached this stage in my life. I would like to thank my mother Dr. Sarita

Gupta and father Dr. Sharad Gupta for their endless support in every difficult situation and

motivating me during my program. I would also like to thank my friends Dr. Anurag

Sikarwar, Nisha Sengar, Dr. Biswajit Chaudhray, Triporna Lahiri, and Subhankar Ghosh here

in Winnipeg who were no less than a family. A very special thanks to Sikarwar and Ghosh

family – and their kids Prisha and Ronav who have given so much love and affection and

made me feel at home always. Also, I would like to thank my overseas childhood friends

Riddima, Ahsu and Vaishali whose precious friendship has given me strength and kept a

smile on my face throughout my life.

I would like to thank my husband Karan Sharma who has been a tremendous support

in this journey, without his unconditional love, patience and positive attitude it would not

have been possible to finish my PhD. Thank you for all the encouragement and motivational

Skype talks that have helped me to be a strong person.

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Table of Contents

Abstract ................................................................................................................................................... ii

Acknowledgements ................................................................................................................................ iv

List of Tables ......................................................................................................................................... ix

List of Figures ........................................................................................................................................ ix

List of Abbreviations ............................................................................................................................. xi

Introduction ............................................................................................................................................. 2

Cardiovascular Diseases and Epidemiology ....................................................................................... 2

Cardiovascular Disease and Cardiac Fibrosis............................................................................... 4

Myocardial Infarction and Wound Healing ........................................................................................ 5

Extracellular Matrix (ECM) .............................................................................................................. 10

Fibrillar Collagens ....................................................................................................................... 10

ED-A Fibronectin .......................................................................................................................... 11

Matricellular Proteins .................................................................................................................. 12

Cardiac Fibroblasts ........................................................................................................................... 13

Origins of Cardiac Fibroblasts: Development and Pathology ..................................................... 14

Cardiac Myofibroblasts ..................................................................................................................... 18

Activation of fibroblast to myofibroblasts – a continuum of phenotype .......................................... 18

Mechanical Stress ......................................................................................................................... 19

Transforming Growth Factor-β .................................................................................................... 20

Persistence of myofibroblasts in the heart following cardiac damage ......................................... 24

Animal Model of MI……………………………………………………………………………………………………………………24

Techniques to study cardiac structure and function………………………………………………………………….25

Cell death mechanisms ..................................................................................................................... 26

Apoptosis ....................................................................................................................................... 26

Necrosis......................................................................................................................................... 27

Autophagy ..................................................................................................................................... 27

Signaling pathways in regulation of autophagy ................................................................................ 33

PI3K-AKT/mTOR signaling .......................................................................................................... 33

Adenosine monophosphate kinase (AMPK) signaling .................................................................. 34

Hypoxia inducible factor-1α ......................................................................................................... 35

Autophagy in Development and Differentiation ............................................................................... 40

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Autophagy in Disease Pathogenesis ................................................................................................. 41

Autophagy in Cardiovascular Diseases ............................................................................................. 42

Autophagy: A therapeutic target in cardiovascular disease .............................................................. 46

Autophagy in phenoconversion or activation of fibroblasts ............................................................. 48

Rationale and Hypothesis…………………………………………………………………………………………………………….50

Rationale…………………………………………………………………………………………………………………………………….51

General Hypothesis ........................................................................................................................ …50

Specific Hypothesis 1. Mechanical stress induces autophagy and activation of cardiac fibroblasts in

vitro ................................................................................................................................................... 52

Objective 1.1: Determine the effect of mechanical stress on cardiac fibroblasts ............................. 52

Specific Hypothesis 2. Autophagy is necessary for mechanical stress-induced activation of cardiac

fibroblasts in vitro ............................................................................................................................. 52

Objective 2.1:Determine the Effect of Inhibition of Autophagy in Cardiac Fibroblasts .................. 52

Objective 2.2: Determine the effect of inhibition of autophagy on myofibroblast phenotype ........... 53

Objective 2.3: Determine the effect of inhibition of autophagy on myofibroblast function .............. 53

Objective 2.4: Determine the effect of inhibition of autophagy on myofibroblast function .............. 53

Specific Hypothesis 3.Inhibition of autophagy in vivo will attenuate cardiac fibrosis following

myocardial infarction ........................................................................................................................ 54

Objective 3.1: Determine the effect of inhibition of autophagy on cardiac remodeling ................... 54

Materials and Methods .......................................................................................................................... 55

Animal Ethics.................................................................................................................................... 55

Cell Isolation and Culture ................................................................................................................. 55

Treatment with Autophagy Inhibitors ........................................................................................... 56

Optimization of Autophagy Inhibitor Treatment by MTT Assay ................................................... 56

Protein Isolation from Adherent Cell Cultures ................................................................................. 57

Western Blot Analysis ...................................................................................................................... 58

Fluorescence Immunocytochemistry ................................................................................................ 59

Scratch Assay .................................................................................................................................... 59

Gel Contraction Assay ...................................................................................................................... 60

Transmission Electron Microscopy (TEM) ...................................................................................... 61

Experimental model of myocardial infarction .................................................................................. 61

Echocardiography ............................................................................................................................. 64

Pathological Analysis of Blood Serum ............................................................................................. 64

Protein Isolation from Whole Tissue ................................................................................................ 64

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Tissue Sectioning for Histology and Immunofluorescence .............................................................. 65

Masson’s Trichrome Staining ........................................................................................................... 65

Statistical analysis………………………………………………………………………………………………………………………..67

Chapter 1: Role of Autophagy in Fibroblast Activation In Vitro .......................................................... 68

Introduction ....................................................................................................................................... 68

Results ............................................................................................................................................... 70

Induction of Autophagy by Mechanical Stress .............................................................................. 70

Inhibition of Autophagy in Cardiac Fibroblasts ........................................................................... 72

Inhibition of Autophagy and Myofibroblast Features ................................................................... 78

Inhibition of Autophagy and Myofibroblast Function .................................................................. 86

Inhibition of Autophagy and Mitogen-Activated Protein Kinase (MAPK) Signaling ................... 93

Discussion ....................................................................................................................................... 101

Chapter 2: Autophagy and Cardiac Fibrosis In Vivo .......................................................................... 108

Introduction ..................................................................................................................................... 108

Results ............................................................................................................................................. 111

Fibroblast phenotype and inhibition of autophagy in the post-MI heart .................................... 111

Assessment of Biochemical Parameters for Cardiac, Liver and Kidney Damage ...................... 131

Functional Assessment Following Myocardial Infarction and Chloroquine Treatment ............ 135

General Discussion ............................................................................................................................. 148

Study Conclusions .............................................................................................................................. 152

Future Directions ................................................................................................................................ 153

Limitations of Study………………………………………………………………………………....156

References ........................................................................................................................................... 157

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List of Tables

Table 1: Types of CVD’s and their pathological features

Table 2: Autophagy proteins and their functions

Table 3: Optimized dosages for autophagy inhibitors

List of Figures

Figure 1: Phases of myocardial wound healing

Figure 2: Sources of cardiac fibroblasts

Figure 3: Activation of cardiac fibroblasts to myofibroblasts

Figure 4: Autophagy pathway and its modulators

Figure 5: Central hypothesis

Figure 6: Study Timeline

Figure 7: Temporal activation of autophagy and fibroblast in P0 adult rat cardiac fibroblasts

Figure 8: Baf-A1, CQ and 3MA treatment inhibits autophagy in P0 cardiac fibroblasts

Figure 9: Ultrastructure of rat ventricular cardiac fibroblasts

Figure 10: Inhibition of autophagy reduces expression of key cardiac myofibroblast markers

Figure 11: Immunofluorescence staining of P0 cardiac fibroblast

Figure 12: Effect of autophagy inhibition on cellular contractility of P0 cardiac fibroblasts

Figure 13: Scratch assay to assess cellular migration of cardiac fibroblasts in the presence and

absence of 7.5 nM Baf-A1, 5mM 3MA, 35 μM CQ

Figure 14: Effect of cell plating and CQ treatment on p38-MAPK phosphorylation in P0

primary cardiac fibroblasts

Figure 15: Effect of MAPK P-38 inhibition on primary rat cardiac fibroblast with and without

CQ treatment

Figure 16: Effect of MAPK inhibition on Baf-A1 treated and untreated P0 cardiac fibroblasts

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Figure 17: Effect of inhibition of autophagy on primary cardiac fibroblast-to-myofibroblast

phenoconversion

Figure 18: LC-3β lipidation as a marker of autophagy in 4 week untreated and chloroquine-

treated post-MI rat hearts (control, myocardium remote to the scar and the infarct

scar).

Figure 19: α-SMA marker levels in 4 week untreated and chloroquine-treated post-MI rat

hearts.

Figure 20: Immunofluorescence staining of post-MI and sham-operated hearts at 4 weeks,

with and without chloroquine treatment

Figure 21: Masson’s trichrome staining for collagen content in 4 week post-infarct left

ventricles, with and without chloroquine treatment.

Figure 22: Hydroxyproline collagen quantification in 4 week post MI cardiac tissue.

Figure 23: LC-3β lipidation as a marker of autophagy autophagy in 8-week untreated and

chloroquine-treated rat hearts post-MI.

Figure24: α-SMA expression in 8-week post-MI hearts treated with chloroquine.

Figure 25: Immunofluorescence staining of 8 week post-MI hearts with and without

chloroquine

Figure 26: Biochemical parameters for cardiac, kidney and liver damage in 4 and 8 week

post-MI rat hearts with and without chloroquine treatment

Figure 27: Ejection Fraction and Fractional Shortening in Post-MI Rats

Figure 28: Heart Rate and Left Ventricular Posterior Wall Diameter in Post-MI rats

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List of Abbreviations

3MA 3-methyladenine

4EBP1 eIF4E-binding protein

ACTA2 Actin-2-α

ADP Adenosine diphosphate

AICAR 5-aminoimidazole-4-carboxamide ribonucleotide

AMP Adenosine monophosphate

AMPK 5' adenosine monophosphate-activated kinase

ANGII Angiotensin II

ANT Adenine nucleotide translocase

α-SMA α-Smooth Muscle Actin

AST/ALT Aspartate transaminase/alanine transaminase

ATP Adenosine triphosphate

Baf-A1 Bafilomycin A1

BAK Bcl-2-homologous antagonist/killer

BAX Bcl-2-like protein 4

Bcl-2 B-cell lymphoma 2

bHLH Basic helix-loop-helix

BMDC Bone marrow-derived cells

BNIP3 B-cell lymphoma 2/adenovirus E1B 19 kDa protein-interacting protein 3

BSA Bovine serum albumin

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CaMKK-b Calcium/calmodulin-dependent kinase kinase-b

CCN CTGF, Cyr61, nov proteins

CCN2/CTGF Connective Tissue Growth Factor

CD34 Cluster of differentiation 34

CD45 Cluster of differentiation 45/lymphocyte common antigen

CK/CK-MB Creatine kinase/creatine kinase-MB

CMA Chaperone-mediated autophagy

COPII Coat protein II

CQ Chloroquine

cTnI Cardiac troponin I

cTnT Cardiac troponin T

CVD Cardiovascular Disease

CyP-D Cyclophilin D

Cyr61 Cysteine rich protein 61

DEPTOR DEP domain-containing mTOR-interacting protein

DMEM Dulbecco's modified Eagle's medium

ECG Echocardiography

ECM Extracellular Matrix

ED-A FN ED-A Fibronectin

EF Ejection fraction

EGTA Ethylene glycol-bis(β-aminoethyl ether-N,N,N',N'-tetraacetic acid)

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eIF4E Eukaryotic initiation factor 4E

EMT Epithelial to mesenchymal transition

EndMT Endothelial to mesenchymal transition

EPDC Epicardium-derived cells

FAK Focal adhesion kinase

FBS Fetal bovine serum

FGF Fibroblast growth factor

FIP200 Focal adhesion kinase family interactin protein of 200 kDa

FN Fibronectin

FoxO3 Forkhead box O3

FS Fractional shortening

GAGs Glucosaminoglycans

GAP GTPase-activating protein

G-CSF Granulocyte colony-stimulating factor

GF Growth factor(s)

GPCR G-protein coupled receptor

GTP Guanosine triphosphate

GβL G protein beta subunit-like

HDAC Histone deacetylase

HIF1α Hypoxia-inducible factor-1α

HR Heart rate

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HRP Horseradish peroxidise

hsc70/hsc40 Heat shock cognate protein of 70/40 kDa

HSCs Hepatic stellate cells

IL Interleukin

IP3 Inositol triphosphate

I-Smad Inhibitory Smad

JNK c-Jun N-terminal kinase

kDa Kilodaltons

kPa Kilopascals

LAD Left anterior descending

LAMP2A Lysosome-associated membrane protein type 2A

LAP Latency-associated protein

LC3 Light chain variant 3

LDH Lactate dehydrogenase

LKB1 Liver kinase B1

LLC Large latent complex

LTBP Latent TGF-β binding protein

LV Left ventricular

LVPWd Left ventricular posterior wall diameter

MAP4K3 Mitogen-activated protein kinase kinase kinase kinase 3

Meox2 Mesenchyme homeobox 2

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MET Mesenchymal to epithelial transition

MI Myocardial Infarction

mLST8 mechanistic target of rapamycin complex subunit LST8

MMP Matrix metalloproteinase

mPTP Mitochondrial permeability transition pore

MRI Magnetic resonance imaging

mRNA Messenger ribonucleic acid

mTOR Mechanistic target of rapamycin

mTORC1/2 Mechanistic target of rapamycin complex ½

NFκB Nuclear factor kappa B

Nov Nephroblastoma overexpressed gene

OPN Osteopontin

P0 Passage 0 (unpassaged)

PBS Phosphate buffered saline

PDGF Platelet-derived Growth Factor

PE Phosphatidyl ethanolamine

PFA Paraformaldehyde

PGs Proteoglycans

PI3K Phosphatidylinositol 3 kinase

PIKK Phosphatidylinositol kinase-related kinase

PINK-1 PTEN-induced putative kinase 1

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PRAS40 Proline-rich Akt substrate of 40 kDa

PRR5 Proline-rich protein 5

PTEN Phosphatase and tensin homolog

PVDF Polyvinylidene difluoride

Rab7 Ras-related GTP-binding protein 7

Rag Recombinant activating gene protein

Raptor Regulatory-associated protein of mTOR

REDD1 Regulated in development and DNA damage responses 1

Rheb Ras homolog enriched in brain

ROS Reactive Oxygen Species

R-Smad Receptor Smad

S100A4 S100 calcium-binding protein A4

S6K1 S6 kinase 1

SAHA Suberoylanilidehydroxamic acid

SDS-PAGE Sodium-dodecyl sulphate polyacrylamide gel electrophoresis

SLC Small latency complex

SMCs Smooth Muscle Cells

S-MEM Minimum essential media Spinner's modification

SMER Small molecule enhancer of rapamycin

Sox9 Sex-determining region Y-box 9

SPARC Secreted protein acidic and rich in cysteine

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TBS Tris-buffered saline

Tbx5 T-box 5

Tcf21 Transcription factor 21

TEM Transmission electron microscopy

TGF-β Transforming Growth Factor-β

TLR Toll-like Receptor

TSP-1 Thrombospondin-1

TTE Transthoracic echocardiography

ULK1 Unc-51 like autophagy activating kinase 1

VDAC-1 Voltage-dependent anion-selective channel 1

Vps34 Vacuolar protein sorting 34

VSMCs Vascular Smooth Muscle Cells

Wk Week(s)

Zeb2 Zinc finger E-box binding homeobox 2

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Introduction

Cardiovascular Diseases and Epidemiology

Cardiovascular disease (CVD) is a category of disease comprising the disorders related to

the heart and the vasculature, involving the circulatory system. CVDs can be described as

conditions resulting in compromised blood supply to and from the heart, causing

cardiovascular dysfunction. As per World Health Organization and Public Health Agency of

Canada guidelines, there are six major types of CVDs. The six categories of CVD are:

Table 1: Types of CVD’s and their pathological features:

No. Types Of CVD Pathological Features

1 Coronary heart disease Complete blockage of one or more coronary

arteries (referred to as MI) leading to tissue

necrosis.

Ischemic heart disease A partial blockage of one or more of the

coronary arteries leading to angina (chest

pain) and dyspnea (shortness of breath).

2 Cerebrovascular disease (stroke)

Blockage of the carotid artery or blood

vessels supplying blood to the brain due to a

hemorrhage (rupture of blood vessel) or

ischemia (blood clots), eventually leading to

stroke.

3 Peripheral vascular disease Reduced blood supply to extremities of the

body such as arms or limbs due to vascular

narrowing, closure, or spasm.

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4 Heart failure

Inability of the heart to pump adequate

blood to the body due to chronic alcohol

consumption, myocardial infarction or

cardiomyopathy.

5 Rheumatic heart disease Bacterial infection in adults, affecting joints,

heart valves, endocardium (endocarditis), or

the pericardium (pericarditis).

6 Congenital heart disease An anatomical birth defect in the heart,

created during fetal cardiac development.

May involve defects of heart valves (such as

being too narrow or complete occlusion);

heart wall defects, such as impaired closure

of the inside walls of the atrium, which can

cause a reverse blood flow to the heart and

lungs; defects of myocardial muscle,

compromising efficiency; or improper

vessel connections, which may improperly

shunt blood travelling through the heart

chambers.

CVDs cause of death, globally representing 31% (17.5 million) of all deaths in 2012 [1].

Also on the global scale, out of 16 million deaths of people under the age of 70 due to non-

communicable disease, 37% are caused by CVDs. Among total CVD-related deaths,

approximately 43% are due to ischemic heart disease, 33% are due to stroke, and the other

26% are due to other causes (congestive heart failure, hypertension, rheumatic fever, etc.)

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[1].Despite remarkable progress in treatment of heart disease, CVD is the leading cause of

death in over one-third of the country’s population [2]. Statistics Canada listed 1.3 million

Canadians living with heart disease in 2011, and around 14,000 died that year due to heart

attack [2].

The Canadian economy incurs $20.9 billion in annual costs due to heart disease and

stroke, even though studies show that 80% of premature heart disease and stroke can be

prevented by adopting a healthy lifestyle [3]. Incidence of cardiac fibrosis, in CVD is

associated with high mortality and there are no effective treatments for its prevention or

reversal post-MI [4].

Cardiovascular Disease and Cardiac Fibrosis

Cardiac fibrosis may be causal to heart failure in CVD [5] and is defined as replacement

and invasion of a healthy functional myocardium by excessive connective tissue if it can

interfere with systolic and diastolic function [6]. Among the six forms of CVD,

coronary/ischemic heart disease has been identified as the most frequent cause of cardiac

fibrosis. Coronary artery disease often leads to MI, which in turn stimulates cardiac fibrosis

[6] and eventual heart failure [5]. Other types of CVD, such as hypertension and diabetic

cardiomyopathy, have also been associated with development of cardiac fibrosis [7-9]. These

pathologies result in damage and loss of the myocardial tissue, ultimately resulting in heart

failure. Two types of cardiac fibrosis have been widely accepted to occur in heart [10, 11],

including:

1. Reactive interstitial fibrosis: This fibrosis is progressive leading to collagen

deposition in interstitium and is seen associated with hypertension, diabetes,

metabolic syndrome and in aging heart. It is an adaptive response to preserve

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cardiac structure and function. There is a diffuse deposition of collagen

throughout the myocardium originating from the microvasculature.

2. Replacement fibrosis (also known as scarring fibrosis): This fibrosis is associated

with MI and cardiotoxicity, among other CVDs. It appears to replace myocytes

lost to necrosis following acute MI [12]. It is localized as in cases of ischemic

cardiomyopathy.

Myocardial Infarction and Wound Healing

In a normal, efficiently working heart there is a constant supply of oxygen and

nutrients provided mainly by the coronary circulation. Approximately 90% of heart attacks

are due to rupture of an atherosclerotic plaque in the coronary arteries, which causes

obstruction of blood supply to the heart via these arteries – also known as ischemia.

Prolonged ischemia leads to myocardial cell death, damage of the cardiac tissue, and loss of

cardiac function. MI can be diagnosed via electrocardiographic (ECG) findings, imaging

techniques such as magnetic resonance imaging (MRI), and by detecting elevated values of

biochemical markers of myocardial necrosis such as cardiac troponin T (cTnT) and I (cTnI)

and creatine kinase-MB (CK-MB) in the blood [13]. With time MI leads to global

pathological remodeling, leading to cardiac hypertrophy and ventricular arrhythmias.

Ultimately, this means that patients who survive acute MI are subsequently susceptible to the

development of heart failure, a leading cause of death [14].

After MI, the injured heart undergoes wound healing following the death of

cardiomyocytes[15]. Rapid necrosis of cardiomyocytes and myocardial tissue loss triggers

three overlapping phases of cardiac wound healing: (1) the inflammatory phase, (2) the cell

proliferation phase, and (3) the ECM synthesis and scar formation phase [16-18].

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1) Inflammatory phase: This acute phase can last for hours or days. Tissue injury causes

release of reactive oxygen species (ROS) and intracellular proteins from the necrotic

cells in the infarcted area, and these products initiate an inflammatory cascade,

including activation of Toll-like receptors (TLRs), the transcription nuclear factor

kappa B (NF-κB), and the complement system [19, 20]. The first cell type to be

recruited after infarction are neutrophils, followed by monocytes [21]. Neutrophils

release inflammatory cytokines such as tumor necrosis factor-α (TNF-α) and

interleukins (IL-3, -6, and -8) that aid in recruiting monocytes, but which can also

lead to further damage and should be controlled [20]. The inflammatory response also

results in activation and release of MMPs (specifically MMP-2 and MMP-9) from

inflammatory cells and fibroblasts that help in ECM degradation [18, 22]. Activation

of the inflammatory phase cleans the dead and damaged tissue which is then followed

by release of TGF-β and IL-10 [23].Both of these factors inhibit inflammation and

induce infarct healing. Healing of the infarct begins with infiltration of monocytes and

lymphocytes[20].

2) Cell proliferation phase: Release of cytokines and growth factors such as TGF-β,

platelet-derived growth factor (PDGF) and fibroblast growth factor (FGF) induces the

proliferation and activation of cardiac fibroblasts to myofibroblasts, for which the

main role is to synthesize new ECM [24].The formation of new vessels to restore

tissue reperfusion (angiogenesis) is also an important feature of this phase [25]. The

closure of the infarct wound occurs via re-epithelization where epithelial epidermal

cells migrate inward to form the wound periphery. Angiogenesis and re-epithelization

help in the formation of granulation tissue, which is the newly formed connective

tissue with blood vessels [18]. Formation of new ECM provides structural integrity,

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and accumulation of ECM components such as fibrillar collagens type I and III,

fibronectin, and glycoprotein signals the final phase of scar formation [26].

3) Extracellular matrix remodeling and scar maturation phase: This phase can continue

for weeks. A shift to matrix synthesis from degradation, due to activated

myofibroblasts in the infarct region, initiates scar formation [18]. With dynamic

changes in ECM structure, interactions between myofibroblasts and the ECM assist

with contracture of the infarct wound. Myofibroblasts form stress fibers containing

contractile proteins such as α-smooth muscle actin (α-SMA) that help in developing

tension needed for correct wound healing [27]. Once the injured tissue is repaired, a

process that can take weeks, resident myofibroblasts are removed via apoptosis [28].

However, this ideal scenario of wound healing might not occur in elderly patients

suffering from other cardiac and non-cardiac ailments. In adverse wound healing, the

myofibroblasts persist in the scar and distal, uninfarcted regions instead of undergoing

apoptosis, and continue to synthesis excess amounts of collagen and ECM proteins,

which lead to stiffening of the tissue, left ventricular dysfunction, and increased

chances of eventual heart failure [29, 30].

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Figure 1: Phases of myocardial wound healing. Post-myocardial infarction (MI) the heart

undergoes overlapping wound healing phases for several days. Cardiomyocyte death occurs

for up to approximately 24 hours, followed by apoptosis or necrosis of the dead tissue,

continuing for up to 2 days post-MI. Thus begins an acute inflammatory phase which

continues for up to 5 days. The hallmark of this phase is recruitment of neutrophils followed

by monocytes and macrophages that phagocytose dead cells. Neutrophils release bioactive

factors such as cytokines, chemokines, and matrix metalloproteinases (MMPs), which help in

ECM degradation and removal of dead tissue. Macrophages recruited by the release of

inflammatory factors from dying myocardial tissue then engulf cellular debris via

phagocytosis. The next phase involves proliferation of cells that will initiate scar formation,

namely fibroblasts and myofibroblasts. The main cell types that initiate this phase are

endothelial cells, which release growth factors (GF), cytokines, and chemokines to induce

angiogenesis and endothelial to mesenchymal transition (EndMT). EndMT causes

proliferation and activation of cardiac fibroblasts to myofibroblasts, and initiates ECM

synthesis. This entire process occurs from 3-28 days. The final phase is scar formation and

9

maturation, which can continue for years. In this phase, activated myofibroblasts release

bioactive factors such as transforming growth factor- β (TGF-β), angiotensin II (ANGII) and

MMPs, which help in accumulation and maturation of ECM proteins. Myofibroblasts also

produce large amounts of ECM structural proteins (primarily fibrillar collagens) and contract,

which helps in wound contracture. In normal wound healing, once the scar tissue is formed at

the site of injury, myofibroblasts undergo apoptosis, halting the process of ECM deposition.

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Extracellular Matrix (ECM)

Cardiac ECM forms the structural scaffold of the heart and is composed of non-

cellular matrix proteins. Cell types present in the ECM include fibroblasts, myocytes,

leukocytes, as well as smooth muscle and endothelial cells of the vasculature. The cardiac

ECM provides mechanical connections between cardiomyocytes, and so is the basis of the

structural and functional integrity of the myocardium [31, 32]. The dynamic nature of the

cardiac ECM means cellular components are under the influence of many cytokines, growth

factors, and hormones that coordinate turnover of matrix proteins and is in constant flux.

Under pathological triggers like ischemia, pressure overload, aging, and infection, the cardiac

ECM can undergo remodeling, primarily due to activation of MMPs [33]. While excessive

production of ECM causes fibrosis and ventricular dysfunction, inadequate ECM deposition

can lead to wall thinning and infarct expansion, resulting in clinical outcomes of aneurysm,

left ventricular (LV) dilatation, and cardiac rupture [34].

The matrix proteins comprising the non-cellular structural aspect of cardiac ECM are

fibrillar collagens, fibronectin, laminin, elastins, glycosaminoglycans (GAGs) and

proteoglycans (PGs). GAGs like hyaluronan, and PGs like syndecans and glypicans (cell

surface proteins) also facilitate with crosstalk between ECM proteins, growth factors, and

cytokines. Hyaluronan is the largest GAG in cardiac ECM and is elevated in MI [35], cardiac

hypertrophy [36] and myocarditis [37]. There are also the non-structural matricellularproteins

that include secreted protein acidic and rich in cysteine (SPARC), thrombospondin (TSP-1),

tenascin-C, osteopontin (OPN), periostin, proteolytic MMPs, and growth factors such as

connective tissue growth factor (CTGF), TGF-β, and PDGF [31].

Fibrillar Collagens

Fibrillar collagen types are the most abundant structural proteins in heart (specifically

collagen type I and III in the interstitium), along with the much less abundant but important

11

non-fibrillar collagen types IV, V, and VI [17]. Studies have shown on days 7 and 21 post-

MI, collagens I and III make up 30% and 60% of infarct scar area, respectively [34].Collagen

synthesis by fibroblasts and myofibroblasts, and other cell types such as vascular smooth

muscle cells (VSMCs) in the ECM, begins with pre-procollagens, which exit the endoplasmic

reticulum and enter the interstitial space. Hydroxylase enzyme processes pre-procollagens

into pro-collagens, transported via vesicles to the Golgi apparatus, where oligosaccharides are

added to form tropo-collagens. Tropo-collagens are linked by the membrane-bound enzyme

lysyl oxidase that acts on tropo-collagen lysine and hydroxylysine residues, resulting in the

formation of collagen fibers. Collagen turnover is critical in the heart, requiring a balance

between synthesis by myofibroblasts and degradation by collagenases (primarily MMP2 and

MMP9) [38]. This homeostasis of ECM is regulated via the TGF-β/Smad signaling pathways

in fibroblasts and myofibroblasts, discussed later [4]. Pathologically, circulating end terminal

peptides of pro-collagen in the serum are biomarkers for myocardial fibrosis, LV dysfunction

and risk of heart failure [34, 39]. Thus, collagen deposition and turnover plays a critical role

not only in scar formation but in overall cardiac remodeling and functioning of the heart post-

MI.

ED-A Fibronectin

Fibronectins (FN) area collection of proteins with a number of subtypes, and in heart

is an ECM glycoprotein expressed by a wide variety of cells and plays a critical role in cell

growth, adhesion, migration, and differentiation [40]. Fibronectins have been shown in

numerous studies to regulate the wound-healing process [41, 42]. Different splice variants

give rise to three modes of fibronectin (types I, II, and III) [43]. Fibronectin Type III

comprises 15 repeating highly conserved segments and has two variable exons extracellular

domain-A (ED-A) and -B (ED-B). The FN type III isoform containing the ED-A splice

variant is termed cellular FN and is present in the cardiac ECM [44]. ED-A FN production is

12

increased with conversion of fibroblasts to myofibroblasts, and is a key molecule involved in

fibrosis in various tissue types, including the lungs, liver, and heart [45]. Under the conditions

of tissue fibrosis, and fibroblast activation, ED-A FN is expressed in abundance by activated

fibroblasts, macrophages, and endothelial cells [32]. Several studies have demonstrated a

direct correlation between increased levels of ED-A FN and cell proliferation, increased cell

adhesion, and activation of fibroblasts to myofibroblasts [46, 47]. ED-A FN-deficient mice

show decreased left ventricular (LV) dysfunction post-MI as compared to wild type mice

[48], suggesting that ED-A FN is critical during developing cardiac fibrosis. Studies have

shown that in the post-MI heart, TGF-β induces myofibroblast secretion of ED-A FN [47, 49,

50]. The exact mechanism for this induction is not known, but Scleraxis, a basic helix-loop-

helix (bHLH) transcription factor, has been identified as the factor regulating TGF-β-induced

fibronectin expression [47, 51].

Matricellular Proteins

Matricellular proteins are the non-structural proteins of the ECM, expressed during

the embryonic development phase and absent in adulthood [52]. Studies have shown that

these proteins are re-expressed during tumor growth and tissue injury to help in matrix

remodeling and wound healing, through regulation of cell proliferation and migration [53].

For example, tenascin-C, important during embryogenesis, is re-expressed by the

mechanically stressed fibroblast specifically near the border zone of the infarct during post-

MI remodeling [54, 55]. Tenascin-C expression can be induced via growth factors such as

FGF and TGF-β, which are released during wound repair. Thrombospondins (TSP) are a

family of secreted glycoproteins that play key roles in cardiac pathologies. TSP-1and TSP-2

are shown to have a protective role in pressure overload (by activating TGF-β to induce ECM

synthesis) and post-MI (via inhibiting MMP2 activation) [31]. Osteopontin (OPN) is another

member of the matricellular proteins, which is up-regulated in macrophages following MI

13

and is important in reducing chamber dilatation, protecting from adverse cardiac remodeling

[56]. Levels of periostin are also increased following cardiac injury. Periostin plays a critical

role in the activation of fibroblasts and subsequent collagen deposition preventing infarct

rupture [31]. However, in a swine model of MI, recombinant periostin delivery to the

myocardium resulted in increased fibrosis [57], the role of this matricellular protein in cardiac

pathology remains unclear.The connective tissue growth factor, Cysteine rich protein 61

(Cyr61), and nephroblastoma overexpressed gene (nov) family of proteins, also known as

CCN proteins, have also been studied extensively in heart disease. Specifically, CTGF has

been elevated post-MI and in the pressure-overloaded heart, and is important in TGF-

signaling, angiogenesis, and myocyte survival [58, 59]. Matricellular proteins can serve as

important diagnostic markers for predicting the extent of adverse ECM remodeling following

myocardial injury. Namely, tenascin-C has been demonstrated as a useful marker for acute

myocarditis, post-MI remodeling, and heart failure; OPN levels serve as a useful marker of

LV dilatation; and serum CTGF correlates with plasma brain natriuretic peptide (BNP) and

TGF-β levels, thus serving as a useful marker for myocardial remodeling [60].

Cardiac Fibroblasts

The main cell types in the heart are cardiomyocytes, which make up 30% of the adult

rat heart, and non-myocytes, which make up the other 70%. Cardiac fibroblasts make up 70%

of this total of non-myocyte cells, which also includes endothelial cells and VSMCs [61-64].

Cardiac fibroblasts are long spindle-shaped cells lacking basement membrane and belonging

to mesenchymal origin [65]. The main function of cardiac fibroblasts is to maintain

homeostasis (synthesis and degradation) of ECM structural proteins. In the normal healthy

heart, cardiac fibroblasts maintain structural integrity through cell-cell and cell-ECM

interactions, ECM homeostasis, proliferation, and migration [62, 66]. Cardiac fibroblasts can

respond to chemical, mechanical, and electrical signals, which can transduce these signals

14

and produce stimulatory factors in response, as a means to signal to other cardiac cell types

like myocytes and endothelial cells to alter their function and affect ECM remodeling.

Fibroblasts secrete a variety of bioactive molecules such as growth factors TGF-B and PDGF,

cytokines such as IL-10, MMPs, and collagens [67]. Fibroblasts play an important role in

maintaining physiological function and affecting cellular function in pathological conditions

[68].

Origins of Cardiac Fibroblasts: Development and Pathology

Cardiac fibroblasts are a heterogeneous cell population as they can originate from

various cell sources under physiologic and pathologic conditions [65]. During embryonic

development, the proepicardial organ has been shown to be the source of the majority of

cardiac fibroblasts. Migratory proepicardial cells give rise to the embryonic epicardium [2].

These cells later undergo epithelial to mesenchymal transition (EMT) to form epicardial

derived cells (EPDCs) that invade atria and ventricle walls and differentiate into cardiac

fibroblasts. EPDC-originating fibroblasts eventually help to form the cardiac interstitium and

annulus fibrosis [69, 70]. EMT is regulated by numerous factors including growth factors

(FGFs, PDGF, and TGF-β) [71, 72], and transcription factors such as Sex-determining region

Y-box 9 (Sox9), T-box 5 (Tbx5), Thymosin β4, and transcription factor 21 (Tcf21) [71-73].

Tcf-21 has been reported to be an essential transcription factor determining that epicardial

cells will differentiate into cardiac fibroblasts [73]. FGF10 has been identified as another

important factor that plays a role in migration of cardiac fibroblast precursors into the

developing myocardium. Studies in FGF10-null mice show that the number of cardiac

fibroblasts in the heart was decreased, and the hearts were smaller in size [74]. Besides acting

as a source of cells of the cardiac interstitium, the endothelial cells in the atrioventricular

canal give rise to valvular fibroblasts, shown by some studies to occur through endothelial to

15

mesenchymal transition (EndMT) under regulation of various cytokines including PDGF,

TGF-β, and Wnt [75, 76].

Both processes (EMT and EndMT) are important drivers of embryonic cardiac

fibroblast maturation and heart development, but can also be activated during adult

cardiopathologies [76]. Proliferation and activation of cardiac fibroblasts is critical to wound

healing, and thus a substantial number of cardiac fibroblasts need to be recruited for proper

cardiac remodeling. Fate mapping studies have shown that EndMT is critical during fibrosis,

with up to 30% of fibroblasts in the post-MI heart originating from endothelial cells [77]. In

pathological scenarios, EMT and EndMT are efficient means for providing a source of

cardiac fibroblasts, which eventually differentiate into myofibroblasts [78]. Under

pathological conditions, profibrotic factors such as TGF-β and hypoxia activate EndMT [79].

Notch signaling has been found to activate the expression of TGF-β and activate EndMT

[80].

It was previously believed that resident fibroblasts were the primary cells contributing

to pathologic remodeling in the injured heart. This suggestion was supported by the

observation that cardiac fibroblasts respond to paracrine and autocrine release of bioactive

molecules such as TGF-β, FGF, and TNF-α released post-injury to initiate wound healing

[76]. In various studies of replacement fibrosis, collagen synthesis and matrix deposition was

initiated by resident fibroblasts at the site of injury [5, 11, 81, 82]. Studies in a pressure-

overload mouse model showed that inhibition of the profibrotic factor TGF-β attenuated

cardiac fibroblast proliferation, ECM production, and myofibroblast activation [82].

However, more recent studies on the origin of fibroblasts in cardiac pathologies have

revealed that the source is not limited to resident cardiac fibroblast, but that there is in fact a

heterogeneous population of cells contributing to cardiac fibroblasts in CVDs.

16

Bone marrow-derived cells (BMDCs) are another important source of fibroblasts in

cardiac pathology, with studies showing that almost 30% of cells in the infarcted area of post-

MI hearts are from a BMDC population in the first 14 days. Around 28 days post-MI, the

percentage of BMDCs in the infarct and border zone was reduced to 17%, with most being

myofibroblasts, as indicated by the expression of myofibroblast marker α-SMA [83]. The

timing of these findings suggests BMDCs play an important role during the early

inflammatory phase of myocardial wound healing. Monocytes also are present in the infarct

scar. Infarcted cardiac tissue shows the presence of cells which co-express monocytic and

myofibroblast markers (S100 calcium-binding protein A4 or S100A4; αSMA) [84].

Additionally, inhibition of monocyte recruitment decreases the cardiac fibroblast population

and myocardial remodeling after myocardial infarction [85].

Circulating fibrocytes are unique fibroblast progenitors of stem cell origin that

express phenotypic similarities with leukocytes. Fibrocytes co-expresses mesenchymal and

hematopoietic markers including lymphocyte common antigen or cluster of differentiation 45

(CD45), cluster of differentiation 34 (CD34), procollagen-1, and vimentin [86]. Cultured

fibrocytes, like myofibroblasts, express α-SMA, which is enhanced by TGF-β1 treatment.

Also like myofibroblasts, cultured fibrocytes exert contractile force which in the heart to aid

in cardiac wound contracture. Fibrocyte activation and trafficking during cardiac repair is a

significant source of myofibroblasts, which are critical to wound healing [87].

Perivascular cells or pericytes have also been studied as a putative source of cardiac

fibroblasts. These cells originate from the perivascular spaces in cardiac vessels [88].

Although pericytes were found to invade the injured kidney in a renal fibrosis model [89], the

role of pericytes in cardiac fibrosis is underexplored.

17

Figure 2: Sources of cardiac fibroblasts. During embryonic development, cardiac

fibroblasts originate via epithelial to mesenchymal transition (EMT)from the pro-epicardium,

or endothelial to mesenchymal transition (EndMT) from the endocardium. In the injured

adult heart, EMT and EndMT embryonic pathways are re-activated to recruit cardiac

fibroblasts for wound repair. In cardiac pathology, sources of fibroblasts also include bone

marrow-derived progenitor cells, monocytes, fibrocytes, and perivascular cells. These various

sources of cardiac fibroblasts also contribute to subsequent fibrosis as a source of

myofibroblast precursors.

18

Cardiac Myofibroblasts

Myofibroblasts were originally identified by Gabbiani et al. as a modified fibroblast

that contributes to growth, development, and tissue repair [27]. These cells share

characteristics with both fibroblasts and smooth muscle cells (SMCs) [27, 90]. Like

fibroblasts, myofibroblasts are a heterogeneous cell type, which can originate from resident

fibroblasts, SMCs, pericytes, and BMDCs (via EMT) [65]. They can produce copious

quantities of growth factors (such as TGF-β, PDGF, and CTGF) and other bioactive

molecules. Compared to fibroblasts, myofibroblast are larger, more motile, and

hypersynthetic for ECM proteins, specifically fibrillar collagens I and III [65].

Myofibroblasts produce stress fibers that coordinate wound contraction through incorporation

of contractile proteins such as α-SMA [91]. This property helps in activation of latent TGF-β,

increased ECM synthesis, collagen fibril reorganization, and development of tensile strength

during the wound healing process [92]. In the healthy heart, myofibroblast are usually absent,

except in valve leaflets. They only appear in the myocardium after cardiac injury in the

resulting scar, which is initially required for reparative fibrosis to prevent LV rupture.

However, their persistent presence and exaggerated fibrotic response is detrimental

eventually, leading to impaired cardiac function and eventual heart failure [93].

Activation of fibroblast to myofibroblasts – a continuum of phenotype

The distinction between cardiac fibroblasts and myofibroblasts is clear-cut with

respect to function, however, the question of how these cells activate in the case of cardiac

pathology has only recently been addressed in the literature. Fibroblast activation is regulated

through various factors, such as mechanical stress, growth factors (primarily TGF-β),

vasoactive peptides (e.g.: ANGII), and cytokines (e.g.: TNF-α)and injury [94]. Cardiac

fibroblasts maintain homeostasis of cardiac ECM, however, with their prolonged activation

mayeventually contribute to cardiac fibrosis [11]. In response to mechanical stress and

19

profibrotic factors, in vitro cardiac fibroblasts are converted to an intermediate phenotype

which is sometimes referred to as the proto-myofibroblast. Proto-myofibroblasts are

distinguished from fibroblasts by the de novo expression of α-SMA and increased expression

of ED-A FN [95]. ED-A FN is one of the ECM proteins expressed in proto-myofibroblasts

and marks the transition to a mature or super-mature myofibroblast [27]. Besides elevated

expression of these markers, mature myofibroblasts have increased focal adhesion proteins

such as vinculin, talin and paxilin [47, 96].

Mechanical Stress

In the healthy hearts, fibroblasts are buffered from exposure to severe mechanical

stress by a stable ECM network [97]. Under normal conditions, fibroblasts are present in an

environment where the external force is between 4 to 18kPa [98]. Due to the

mechanosensitive nature of cardiac fibroblasts, the first stimulus that activates a phenotypic

shift of fibroblast to myofibroblast is the change in mechanical environment following injury.

Architectural integrity of the tissue is lost and resident fibroblasts are then exposed to

constant mechanical stress. In the fibrosing heart, ECM stiffness is increased to around

100kPa [99]. Cardiac fibroblasts interact with the ECM through transmembrane proteins

called integrins, which translate these mechanical signals into the intracellular response,

including activation of profibrotic signaling cascades including the TGF-β/Smad and MAPK

pathways.

Constant mechanical stress and stiffened ECM results in mechanical activation of

latent TGF-β in the cardiac ECM, which induce a vicious cycle for secretion of TGF-β that

also stimulates further collagen production [100]. Mechanical tension has been shown to

induce ANGII production, which can further enhance TGF-βsignaling. ED-A FN expression

is also upregulated by increased mechanical tension [47]. McCulloch et al. examined the

effect of force-induced α-SMA expression in rat cardiac fibroblasts, and found that culture of

20

cardiac fibroblasts on rigid substrate increases cell tensile force and is associated with

increased α-SMA expression, both suggesting conversion to the myofibroblast phenotype. A

study on human burn scar biopsies showed that when scar tissue is subjected to mechanical

tension, a high number of α-SMA positive myofibroblasts are observed in the injury

site[101]. An in vivo study has also shown that mechanical tension in tissues is correlated

with increased expression of α-SMA [101]. Expression of myofibroblast markers α-SMA and

ED-A FN, and fibroblast activation to the profibrotic myofibroblast phenotype is

mechanically regulated [102, 103]. These changes results in excessive ECM deposition and

distorted architecture, which eventually causes pathological remodeling [5].

Transforming Growth Factor-β

Cardiac fibroblasts express various receptors on their cell surface which are

associated with fibrosis such as receptors for TGF-β, ANGII and PDGF. The ANGII type 1

receptor (AT-1) can induce increased mRNA and protein levels of TGF-β, and vice versa. It

has been shown in a study, that rat cardiac fibroblasts stimulated with TGF-β responded with

a significant increase in α-SMA and collagens type I and III, as compared to untreated

controls [104]. TGF-β, in an autocrine fashion increases the constitutive expression of α-

SMA stress fibres in the activating myofibroblast, and also upregulates its own mRNA and

protein expression [101]. Both mechanical tension and stimulation by TGF-β are necessary

for developing α-SMA positive myofibroblasts [105, 106]. Profibrotic TGF-β is the primary

growth factor involved in activation of myofibroblasts in various tissues including the

myocardium, and the development and maintenance of cardiac fibrosis.

Initially, TGF-β is released into the cardiac interstitium as an inactive dimeric form by

numerous cell types including cardiac fibroblasts and myofibroblasts. This inactive form,

latent TGF-β, is bound in the ECM until activated by mechanical stress and other profibrotic

factors. The TGF-βgene produces an N-terminal region called latency associated protein

21

(LAP) that non-covalently binds with TGF-β and forms a small latency complex (SLC). Once

released into the ECM the SLC forms a disulfide linkage with the large latent TGF-β-binding

protein (LTBP) to form a large latency complex (LLC). Further activation of the latent TGF-

β requires cleavage of LAP from this complex. Various factors can induce this cleavage such

as proteases (MMPs, TSP-1, and plasmin), acidic pH, release of ROS, or mechanical stress

[101, 107]. Myofibroblast contraction mediated by β1, β3, and αvβ5 integrin binding to LAP

can directly activate latent TGF-β1 from LLC in a protease-independent manner [106].

Once activated, TGF-β1 is released from the LLC into the ECM, where it binds to its

receptor, the serine threonine kinase cell surface homodimer TβRII [105]. TβRII then

undergoes autophosphorylation and activates TβRI forming a heterotetrameric complex

[108]. Intracellular receptor Smads (R-Smads) Smad2 and Smad 3 form a complex

(Smad2/3) that binds an intracellular phosphorylation site on TβRI. This initiates a cascade of

Smad-dependent signaling, resulting in formation of a downstream Smad3 and Co-Smad

4complex. The Smad3/4 complex translocates to the nucleus and upregulates the transcription

of fibrotic gene such as collagen I&III [109]. In the post-MI heart, there is a significant

increase in R-Smad2/3 phosphorylation coinciding with nuclear Smad accumulation and

increased collagen deposition, suggesting a critical role of TGF-β/Smad signaling in

transcriptional activation of profibrotic genes such as collagen and α-SMA.

Conversely a group of Smads inhibit gene expression called I-Smads, which includes

Smad6 and -7.I-Smad 7 inhibits TGF-β/Smad signaling by disrupting activation of R-Smads

by the TGF-β1 receptor [110]. The nuclear protein Ski, which appears to regulate anti-fibrotic

effects in fibroblast to myofibroblast conversion, is another negative regulator of TGF-β,

acting as an endogenous Smad inhibitor. Ski represses Smad-mediated TGF-β signaling by

inhibiting activation of profibrotic gene promoters by Co-Smad4 in the nucleus [111, 112].

However, Ski cannot act as a transcriptional inhibitor via direct binding to DNA, but requires

22

the presence of the Smad complex [112]. Ski is also assisted by other transcriptional regulator

proteins such as Meox1, Meox2, and Zinc finger E-box-binding homeobox 2 (Zeb2). Meox1

and Meox2 are homeodomain proteins required in development of murine skeletal muscle

[113]. Both are important in fibrosis. Meox2 can be down-regulated by physical stimuli, such

as mechanically damaged arteries [114]. Anti-fibrotic transcription factor Meox2 has been

found to inhibit profibrotic TGF-β effects, specifically by inhibiting EMT [115]. Zinc finger

E-box binding homeobox 2 (Zeb2), known for its role as a co-repressor in Smad-mediated

TGF-β signaling, has been repressed by nuclear Ski over-expression. Zeb2 repression of

mesenchyme homeobox 2 (Meox2) is relieved, resulting in upregulation of Meox2. Increased

Meox2 expression inhibits activation of profibrotic genes includingα-SMA, ED-A FN, and

collagens. However, in the injured heart, Ski is translocated to the cytosol, allowing Zeb2 to

down-regulate Meox2 – inducing expression of profibrotic genes and fibroblast to

myofibroblast phenotype conversion. This mechanism shows how Ski plays a role as

antifibrotic protein during ECM remodeling post-MI [116].

23

Figure 3: Activation of cardiac fibroblasts to myofibroblasts. In the healthy heart,

extracellular matrix (ECM) components are continuously turned over through synthesis and

degradation by interstitial cardiac fibroblasts. In cardiac fibrosis, myofibroblasts play an

important role in excessive synthesis and remodeling of the cardiac extracellular matrix in

response to mechanical stress or stimulation by profibrotic factors such as transforming

growth factor-β1 (TGF-β1) and angiotensin-II (ANGII). In these pathologies, adult

fibroblasts undergo a phenotypic transition to myofibroblastic cells, and thus acquire

contractile and hypersecretory characteristics. This transition is marked by increased α-SMA,

ED-A FN, and collagens type I and III.

24

Persistence of myofibroblasts in the heart following cardiac damage

Fibroblasts migrate to the infarct zone in post-MI hearts and undergo activation to

myofibroblasts and repopulate the infarct zone after the loss of cardiomyocytes [117].

Although myofibroblasts are necessary for wound healing after ischemic injury, their

persistent presence and excess production of collagen after proper wound healing leads to the

development of cardiac fibrosis, resulting in a stiff and non-compliant heart. Persistence of

myofibroblasts in the infarct and distal regions of the post-MI heart is purportedly caused by

a failure of these cells to undergo regulated cell death through apoptosis or other mechanisms

[118]. One potential cause of this failure of myofibroblasts to cease wound healing activities

is ongoing neuroendocrine dysregulation, potentially due to high wall stress resulting from

the presence of collagen-rich scar tissue. This environment may confer intermittent bouts of

ischemia, and therefore heart remains in a constant wound healing state, which contributes to

activated and persistent myofibroblasts in the myocardium [119].

Animal Models of MI:

Rat models have dominated research in cardiac biology due to many benefits over of

mice such as low cost, ease of handling and their larger size. In heart failure models,

myocardial injury is induced by various procedures: surgical, pharmacological, or

cauterization. The surgical method was first developed by Hans Selye and modified by

Pfeffer and coworkers, where they ligated the left coronary artery [120][121]. The procedure

requires performing a left thoracotomy on the anesthetized rat. Gentle pressure is applied on

the right side of the thorax and the heart is exteriorized. The left coronary artery is ligated

between the pulmonary artery outflow tract and the left atrium and the heart is returned to its

normal position. Finally the thorax is immediately sutured and closed [122]. Left coronary

artery ligation is the most common method used to induce acute myocardial damage in rat

and other animal models. Another widely used experimental model for induction of infarction

25

is achieved by administration of isoproterenol [123]. Isoproterenol is a beta-one adrenergic

receptor (B-AR) agonist and at high doses it induces cardiac myocyte necrosis and extensive

LV dilatation and hypertrophy [124]. The third method which is not used frequently due to

the limitations of reproducibility is the cauterization method. It consists of generating

overlapping burns in exposed rat hearts by applying a 2-mm-tipped soldering iron to the

epicardium of the left ventricle [125]. This where the animals show induction of transmural

injury in the left ventricle, with a low incidence of postoperative mortality [126].

Techniques used to study post-MI cardiac structure and function

Hemodynamic studies: The post–MI rat model is subjected to an invasive technique

where an atrial catheter is used to obtain systemic and LV blood pressure from

femoral and carotid arteries, respectively. Hemodynamic studies provide information

about systolic, diastolic and mean arterial pressure (MAP); heart rate (HR) as well as

central venous pressure (CVP) [127]. In post-MI rat model a reduction in load

dependent parameters of cardiac function (e.g. SV (stroke volumes), CO (Cardiac

Output), EF, dP/dt max/min) is observed at the end of 4 weeks, compared to sham-

operated control animals. These parameters provide an evidence of structural

changes in the myocardium such as scar thinning and myocardial hypertrophy at 4

weeks post MI [128].

Echocardiography: Echocardiography is one of the most useful diagnostic tools used

for assessing parameters of cardiac remodeling in small animal models of MI [129].

Along with the advantage of being non-invasive it is reproducible and gives accurate

estimations of the cardiac structure and function [130]. M-mode echocardiography is

widely used in laboratory experimental animals to measure the dimensions of cardiac

chambers and cardiac wall motion abnormalities [131]. It can also measure HR, SV,

LVESV, LVEDV, wall thickness (LVPWd), EF % and E/A ratio (ratio of peak

26

velocity flow in early diastole (the E wave) to peak velocity flow in late diastole

caused by atrial contraction the A wave ). M-mode echocardiography has a high

temporal and spatial resolution [132].

Magnetic resonance imaging (MRI): MRI is a very efficient and non-invasive

technique used to study the changes in left-ventricular geometry and function post-MI

[133] This technique can yield information on the myocardial function, left

ventricular mass and wall thickness in various animal models [134]. MRI also

provides valuable information about cardiac remodeling post-MI without the need to

sacrifice animals. Assessment of infarct size is an important parameter when studying

LV remodeling. This is possible with the MRI and delayed enhancement imaging

technique, where a contrasting agent such as gadolinium is administered and it allows

myocardial viability and infarct size to be determined [136]. Futhermore, the

advantages of MRI include its tomographic nature with high spatial and temporal

resolution and excellent soft tissue contrast [135].

Cell death mechanisms

Cell death mechanisms not only play an important role in homeostasis for

physiological function, but are also important in disease pathogenesis. The three main

mechanisms of cell death are apoptosis, necrosis, and autophagy. The primary mechanisms of

cell death are apoptosis and necrosis, which are distinctly regulated but share some

overlapping features.

Apoptosis

Apoptosis is characterized by cell shrinkage, cell fragmentation into membrane

enclosed apoptotic bodies, and phagocytosis by nearby cells, which acts as a clean-up

operation to prevent inflammation [137]. Apoptosis has been shown to be mediated by

27

intrinsic and extrinsic pathways and is dependent on caspases [138]. Cellular ATP levels are

maintained via continuous production and low energy expenditure.

Necrosis

Necrosis is characterized by loss of cell membrane integrity, cellular and organelle

swelling, and the onset of inflammation. It releases pro-inflammatory factors that activate the

inflammasome, further perpetuating the release of pro-inflammatory interleukins [139].

Unlike in apoptosis, adenosine triphosphate (ATP) levels and energy expenditure are

dramatically reduced in necrotic cells, due to severe mitochondrial damage [140].

Autophagy

Autophagy is a catabolic quality control mechanism for turnover of long-lived proteins

and organelles in bulk through lysosomal degradation. Autophagy occurs constitutively at

low levels in eukaryotic cells to perform housekeeping functions (eg, the destruction of

dysfunctional organelles). Breakthroughs in the field of autophagy research have come from

studies in yeast (Saccharomyces cerevisiae), including the discovery of over 32 autophagy-

related Atg genes [141]. Autophagy can be up-regulated by external stressors such as

starvation, oxidative stress, or hormonal imbalance, as well as in response to internal needs

such as removal of protein aggregates. This indicates that the process of autophagy is an

important survival mechanism [142]. To date three types of autophagy have been identified:

1. Chaperone-mediated autophagy (CMA): Chaperone molecules selectively target

cytosolic or membrane proteins containing the specific amino acid KFERQ motif for

lysosomal degradation. It is unique in its selectivity and direct shuttling of components

to the lysosome. In CMA, the chaperone heat shock cognate protein of 70 kDa (hsc70)

recognizes the specific KFERQ motif present on various proteins and facilitates their

interaction with lysosomes through lysosome-associated membrane protein type 2A

28

(LAMP2A)which acts as the receptor for the hsc70-target protein complex. Additional

chaperone proteins at the lysosomal membrane (Bag1, hip, hop, and hsc40) facilitate

unfolding of the target protein, and LAMP2A multimers form a translocation complex

that allows transport of the target protein to the lysosome [143]. CMA is increased with

elevated oxidative stress in the cell and causes degradation of oxidized proteins, acting

as an important adaptive cellular response to stress conditions [144].

2. Microautophaghy: Originally observed in yeast, microautophagy is a generally non-

selective process that involves cytosolic contents being directly internalised by the

lysosomal vacuolar membrane. It is especially important in cellular survival in

response to starvation, nitrogen deprivation, or rapamycin treatment. Unfortunately,

the molecular mechanisms that mediate microautophagy and its function in

mammalian cell physiology and pathology remains elusive [145].

3. Macro-autophagy –This is the most prevalent form of autophagy (hereafter simply

referred to as autophagy). In this process the whole organelle and misfolded proteins

are engulfed by a double membrane structure to form the autophagosome, which then

fuses with lysosomes to form autophagolysosomes [146, 147].

Autophagy is divided into four major steps. These are nucleation, elongation, closure, and

maturation, as outlined:

1. Nucleation: Also termed the initiation step or membrane isolation step, nucleation and

assembly of the initial phagophore (also called an isolation membrane) is initiated

usually by mechanistic target of rapamycin (mTOR) repression in response to nutrient

deprivation. One of the initial steps of nucleation involves activation of the class III

phosphatidylinositol 3-kinase (PI3K) vacuolar protein sorting 34 (Vps34). This

requires formation of a multiprotein complex composed of Vps34, a myristoylated

serine/threonine kinase p150, Atg14, and Beclin-1 [148]. Formation of this complex

29

induces formation of the phagophore [149, 150]. This step can be inhibited chemically

with 3-methyladenine [151].

2. Elongation: Involves the formation of a double-membrane vesicle structure through

two evolutionarily conserved (in both yeast and mammals) ubiquitin-like conjugation

systems. The first mechanism requires covalent conjugation of Atg12 to Atg5,

facilitated by E1-like Atg7 and E2-like Atg10, which is organized into a complex with

Atg16 (the Atg5-12-16 complex). The second mechanism involves

phosphatidylethanolamine (PE) conjugation with LC3 through protease action of

Atg4, Atg7, and E2-like Atg3. Proteolytic cleavage of the C-terminal region of LC3

by Atg4 converts it to LC-3I. LC-3I further conjugates with phosphatidyl

ethanolamine (PE) with the help of two ubiquitin-like modifier-activating (E-like)

enzymes Atg7 and Atg3. This lipid conjugation converts soluble LC3 to LC-3II (or

LC3-PE), the autophagy vesicle-associated form[152]. The fusion of this complex

with the membrane results in vesicle elongation.

3. Closure: Lipidated LC3 (eg,: LC-3II) and the associatedAtg5-12-16 complex

facilitatethe closure of the autophagosome and interact with an adapter protein p62.

p62 directly binds both poly- or mono-ubiquitinated proteins via its ubiquitin-

associated (UBA) domain [153, 154], and links the ubiquitinated cargos to the

autophagy machinery for autophagic degradation.LC3-II is the ‘gold standard’ used to

measure autophagy flux (i.e., the rate of formation of autophagosomes)[155-157].

4. Maturation: After phagosome formation, they undergo maturation through lysosomal

fusion forming autolysosomes. This final step of autophagy is governed by

translocation and fusion of autophagosomes with lysosomes. In yeast, maturation

occurs with the assistance of multiple proteins like SNAREs and UVRAG to form

autolysosomes [158]. In mammals, this process is not well understood, but involves

30

LAMP-2 and the GTP-binding protein Ras-related GTP-binding protein 7 (Rab7)

[159, 160]. Autolysosomes sequester damaged proteins and organelles, LC3 is

cleaved off by Atg4, recycled with other components, and subsequently degraded

such that its building blocks can be reused (see Figure 4) [161].

Apart from the role of Atg proteins, the cellular cytoskeletal system and secretory

pathways are also important in autophagy. Certain guanosine triphosphate (GTP) exchange

factors (such as Sec12 and Sec16,and two coatomer subunits of the coat protein II (COPII)

coat, Sec23 and Sec24) are needed for the biogenesis of autophagosomes [162]. Microtubules

are involved in autophagy in higher eukaryotes, as evidenced by inhibition of autophagosome

formation with the microtubule depolymerizing drug nocodazole in rat hepatocytes[163].In

mammalian cells, autophagosomes are formed at random locations in the cell and transported

toward the nucleus by the motor protein dynein, which requires the tubulin deacetylase

histone deacetylase 6 (HDAC6) for autophagic clearance of cellular components [164, 165].

31

Figure 4: Autophagy pathways and its modulators. Various growth and nutrient signaling

pathways are associated with regulation of autophagy. Following inhibition of mTOR, the

ULK/Atg13/FIP200 complex is activated and initiates autophagosome/phagophore formation.

The class III PI3 kinase (Vps34)-Atg14L-Becn1 complex also regulates the autophagosome

nucleation step. To expand the autophagosome membrane, two ubiquitin-like conjugation

systems are required for conjugation of LC3 and Atg12 to phosphatidyl ethanolamine (PE) on

the autophagosome membrane and Atg5, respectively. Further, the Atg12-Atg5 conjugate

interacts with Atg16, presumably located on the surface of the autophagosome membrane.

The complete autophagosome fuses with the lysosome to form the autolysosome, and cargo

molecules engulfed by autophagosomes are degraded by lysosomal hydrolases and recycled

back to the cytoplasm. Several pharmacological inhibitors (red) modulate distinct steps of

autophagy. Some autophagy proteins with enzymatic activity (shown in green and yellow)

32

could be crucial target proteins for modulation of autophagy. Points where inhibitors could be

potentially developed are shown as blank boxes. CQ, chloroquine. Used with permission from

Heesun C et al., Nature Biotechnology, 2012.

33

Signaling pathways in regulation of autophagy

PI3K-AKT/mTOR signaling

Growth factors, G-protein coupled receptors (GPCR) and tyrosine kinase receptors on

the membrane activate the PI3K/Akt signaling pathway which induces cell growth,

differentiation and survival. PI3K/Akt signaling activates mTOR, which is a negative

regulator of autophagy and thus promotes survival [166]. mTOR is an evolutionarily

conserved serine/threonine protein kinase belonging to class I of the PI3K family that

regulates cell growth and metabolism as well as regulating autophagy [167-169]. Two

functionally distinct complexes are formed by mTOR: the autophagic regulator mTOR

complex 1 (mTORC1), and cell metabolism and cytoskeleton regulator mTOR complex 2

(mTORC2) [168]. In response to the nutrient environment, mTORC1 either down-regulates

autophagy (nutrient-rich environment) or upregulates autophagy (nutrient

deprivation).mTORC1 is composed of mTOR, regulatory-associated protein of mTOR

(Raptor), G protein beta subunit-like (GβL), and DEP domain-containing mTOR-interacting

protein (DEPTOR). This complex is directly inhibited byrapamycin [170]. During nutrient

deprivation, mTORC1 function is inhibited, allowing activation of autophagy through the

PI3K-adenosine monophosphate-activated protein kinase (AMPK)/mTOR axis [171, 172]. In

mammals, mTOR/Raptor-dependent signalling is mediated through amino acids via

activation of class III PI3K, while growth factors mediate this action through class I PI3K

[173, 174].

Amino acid activation acts downstream of phosphatases and endocytic trafficking

regulated by upstream mTORC1 kinase, Ste-20-related mitogen-activated protein kinase

kinasekinasekinase 3 (MAP4K3) and Vps34 [170, 175]. In addition, the GTPases

recombinant activating gene protein (Rag) and Ras homolog enriched in brain (Rheb) are also

important amino acid-dependent regulators of mTORC1. Amino acid-rich conditions favour

34

the active conformation of the Rag heterodimer (RagA/BGTP

-RagC/DGDP

) which interacts

with Raptor to translocate mTORC1 to the lysosome [176, 177]. The Rag heterodimer also

interacts with LAMP2 and Rheb, promoting further activation of the mTORC1 signaling

pathway and repression of autophagy. The tuberous sclerosis complex (TSC) proteins 1 and 2

(TSC1 and TSC2)serve as a checkpoint for autophagy by forming a TSC1/2 complex to

inhibit Rheb, decreasing mTORC1 activity and resulting in activation of autophagy [169,

171]. However, PI3K signaling will inhibit TSC1/2 by phosphorylation, allowing Rheb

activation, mTORC1 activity, and repression of the autophagy process [178].

The role of mTORC2, a complex comprising mTOR, GβL, rapamycin-insensitive

companion of mammalian target of rapamycin (RICTOR), and mSIN1, is less understood in

autophagy. Unlike mTORC1, mTORC2 is not directly inhibited by rapamycin.mTORC2acts

as a negative feedback activator and helps in inducing autophagy. mTORC2 is directed

towards Akt through action of the RICTOR subunit, and further to forkhead box O3 (FoxO3),

which is blocked by Akt [179, 180]. PI3K/pAKT does not regulate autophagy only via

mTORC-1; but through FoxO3 mediated transcriptional regulation of two autophagy related

genes LC3 and Bnip3 (a BH3-only protein), leading to the induction of autophagosome

formation [181].

Adenosine monophosphate kinase (AMPK) signaling

AMPK is an important regulator of autophagy. Under nutrient- and energy-deprived

conditions the ratio of adenosine monophosphate (AMP)to adenosine diphosphate (ADP)

increases, which activates AMPK viaphosphorylation by liver kinase B1 (LKB1) [182] and

calcium/calmodulin-dependent kinase kinase-b (CaMKK-b) [183]. Activated AMPK acts as

an upstream regulator of mTOR-1 and initiates autophagy by repressingTSC2 activity [184].

Activated AMPK phosphorylates elongation factor-2 kinase, balancing peptide elongation

and autophagy [185]. The unc-51 like autophagy activating kinase 1 (ULK1) complex

35

comprises Atg13, focal adhesion kinase (FAK) family interacting protein of 200 kDa

(FIP200) and Atg101 [186], which is phosphorylated by the AMPK/mTOR energy sensing

pathway and represses autophagy induction [187].

Hypoxia inducible factor-1α

Under conditions of hypoxia and oxidative stress the hypoxia-inducible factor-1α

(HIF1α) transcriptional factor accumulates in the nucleus and controls expression of various

genes that are important to metabolic adaptation [188]. Two important genes induced under

hypoxia are B-cell lymphoma-2/adenovirus E1B 19 kDa protein-interacting protein 3

(BNIP3) and BNIP3L. Protein products of there genes have been shown to increase

mitochondrial autophagy and thus reduce ROS production [189, 190]. BNIP3 and the protein

regulated in development and DNA damage responses 1 (REDD1) negatively regulate

mTORC1 activation through binding of the small GTPase Rheb and activation of the TSC

complex, respectively [191, 192]. Additionally, BNIP3 competes with Beclin-1 for B-cell

lymphoma2 (Bcl-2) binding, thus promoting autophagosome formation. The proteins

involved in autophagy and their functions are found in Table 1.

36

Table 2: Autophagy proteins and their functions.

Adapted from Ghavami. S, Gupta. S, et al. Autophagy and heart disease: implications for

cardiac ischemia-reperfusion damage, Current molecular medicine, 2013.

Protein Function Reference

Akt/protein

kinase B

Serine/threonine kinase protein; regulates glucose metabolism

(Akt2), cell growth, migration, and inhibition of apoptosis

(Akt1). Interacts with and is activated by class I PI3Ks.

Downstream of Akt activation, can further stimulate mTORC;

suppresses autophagy in PI3K-independent manner; can directly

phosphorylate Beclin-1 and inhibit formation of autophagy

initiation complex, suppressing autophagy

[193]

Atg 5-12-16

complex

Atg 5-12-16 complex, assisted by Atg10 and Atg7; during the

formation of autophagosomes, gets localized at the membrane

[25]

Atg14 Cleaves C-terminus from LC-3, yielding cytosolic LC-3I [25]

Atg7

Atg 7 is a homodimeric E1 enzyme. Interaction of Atg 7 with

Atg3 assists in conjugation of LC-3I and its interaction with Atg

10 helps in formation of Atg 5-12 complex, during initiation of

autophagy.

[148]

Beclin-1 Initiates formation of phagophore along with Vps34 and Atg14. [25]

Bnip3

B-cell lymphoma 2/adenovirus E1B 19 kDa protein-interacting

protein 3 (Bnip3); belongs to pro-apoptotic B-cell lymphoma 2

(Bcl-2) family of proteins; can regulate various cell death

pathways (apoptosis, autophagy, and necrosis); activates

autophagy in hypoxia via opening of mPTP in mitochondria

[194]

37

FoxO3

Forkhead box O3 (FoxO3); transcription factor characterized by

forkhead DNA-binding domain; induces autophagy in skeletal

muscle atrophy; regulates induction of Atg’s.

[181]

HIF1

Hypoxia-inducible factor 1 (HIF1); transcription factor

belonging to basic helix-loop-helix (bHLH) family; a

heterodimer composed of HIF-1β subunit and an oxygen-

regulated HIF-1α subunit; expressed in response to hypoxia for

cell survival; under hypoxic stress can induce autophagy and

cause epithelial to mesenchymal transition (EMT) in cancer

cells; can also activated Bnip3 inducing mitophagy via

disruption of Bcl-2/Beclin-1 interaction, releasing Beclin-1 to

initiate autophagy.

[190]

LC-3 β

Atg8 (in S. cerevisiae)/LC-3 β (in mammals); light chain 3

microtubule-associated protein. Cleavage at C-terminal yields

LC-3I; LC-3I lipidation yields LC-3II; LC-3II associates with

double membrane of autophagosome on luminal and

cytoplasmic phase. Used as marker of autophagy

[25]

mPTP

Mitochondrial permeability transition pore (mPTP); composed

of voltage-dependent anion channel (VDAC), adenine

nucleotide translocase (ANT), and cyclophilin D (CyP-D);

production of reactive oxygen species leads to opening of mPTP

and subsequent membrane depolarization, further leading to

autophagy of damaged mitochondria (mitoautophagy)

[62]

mTOR

Mammalian target of rapamycin (mTOR); a serine/threonine

protein kinase of large size (near 300 kDa); belongs to

[62]

38

phosphatidylinositol kinase-related kinase (PIKK) family.

Activity of mTOR is inhibited under nutrient starvation, a

critical step for autophagy induction in eukaryotes

mTORC1

mTOR complex 1; contains Raptor (KOG1 ortholog),

GβL/mLst8, proline-rich Akt substrate of 40 kDa (PRAS40) and

DEP domain-containing mTOR-interacting protein (DEPTOR);

key regulators of translation and ribosome biogenesis in yeast

and mammalian cells (respectively); responsible for autophagy

induction in response to starvation.

[62]

mTORC2

mTOR complex 2; contains Rictor (Avo3 ortholog), GβL/mLst8,

Sin1 (Avo1 ortholog), proline-rich protein 5 (PRR5/protor), and

DEPTOR. Rictor is a rapamycin insensitive companion of the

mTORC2 complex; mTORC2 is involved in regulating the

phosphorylation and activation of Akt/protein kinase B and

protein kinase C.

[62]

p62

Ubiquitin-binding scaffold protein; acts as cargo for localization

of ubiquitinated protein aggregates to autophagosomes for

degradation. Accumulates with inhibition of autophagy; used as

marker of autophagic flux.

[66]

PI3K

Signal transducer enzymes; phosphorylate the third hydroxyl

group of inositol ring of membrane-bound phosphatidylinositols.

Four major classes, based on structure: class I, II, III, and IV.

Class I substrates important for downstream Akt/mTOR

signaling

[195]

Raptor Regulatory associated protein of mTOR1 complex; an [180]

39

evolutionarily conserved adaptor protein of mTORC1; regulates

mTORC pathway by interacting with mTOR and its substrates,

including ribosomal protein S6 kinase 1 (S6K1) and eukaryotic

initiation factor 4E (eIF4E)-binding protein (4EBP1)

Rag and

Rheb

Recombinant activating gene (Rag) and Ras homolog enriched

in brain (Rheb) are GTP-binding proteins regulating mTORC1

activity; in presence of amino acids, Rag GTPases localize

mTORC1 to lysosomes; Rheb stimulates mTORC1 kinase

activity via direct interaction or conformational change,

eventually leading to enhanced substrate phosphorylation and

mTOR pathway activation

[196]

Rictor

Rapamycin-insensitive companion of mammalian target of

rapamycin (Rictor); a component of the mTORC2 complex;

unlike Raptor, is resistant to rapamycin treatment. Rictor and

mTOR2 activation leads to actin organization during embryonic

development; in conditions of excessive autophagy induction,

promotes activation of mTOR pathway and halts autophagy.

[197]

TSC1/2

complex

Tuberous sclerosis 1/2 (TSC1/2) complex, belonging to the

tumour suppressor genes; plays a major role in mTOR signaling;

GTPase-activating protein (GAP); inhibits activity of Rheb by

converting Rheb-GTP to Rheb-GDP; acts as negative regulator

of mTORC1 activation.

[198]

Vps34

Class III phosphatidylinositol 3 kinase is crucial for phagophore

formation and elongation. It also assist recruitment of other

ATG proteins

[23]

40

Autophagy in Development and Differentiation

Autophagy is a dynamic process important in cell death and contributes to

development and differentiation [199]. During oocyte preimplantation, oocytes undergo

autophagy four hours post-fertilization in response to calcium oscillations wherein autophagy

is important for activating transcription and also for elimination of excess maternal proteins

[200, 201]. During the neonatal stage all tissues except the brain exhibit autophagy because

of starvation, as the maternal supply of nutrients from the placenta is ended. In this stage,

autophagy allows amino acids and other energy resources to be provided to the neonate

through activation of low energy-sensing AMPK and subsequent induction of various Atg

proteins [202]. The critical role of autophagy in development is evidenced by embryonic

lethality caused by Atg5 mutation and neonatal lethality in response to mutations of Atg3, 7,

9, and 16 [199]. Autophagy also plays an important role in cell differentiation, including

erythrocyte development, for removal of mitochondrial and other sub-cellular organelles

[203], in T and B lymphocyte differentiation [204], and in differentiation of adipocytes [205].

In terminally differentiated cells such as neurons and cardiac cells, autophagy processes

maintain cellular homeostasis. In the cardiovascular system, an optimal level of autophagy

has been found to be is critical within the following cell types- endothelial cells, vascular

smooth muscle cells, fibroblast, macrophages, and in cells of the myocardium. Excessive or

insufficient autophagy within these cell types can contribute to cardiac pathologies [206].

The zebrafish has been a model organism for most of the work demonstrating the role

of autophagy in cardiac development, with very few reports in the hearts of developing mice.

Studies in zebrafish demonstrated that morpholino and Atg mutations result in defective

cardiac and vascular development [207]. FGF is an important growth factor in cardiac

development which regulates cardiac stem cell differentiation to cardiomyocytes. In the

41

presence of FGF, autophagy is maintained at a low level whereas in its absence, autophagy is

highly induced, resulting in induction of cardiomyocyte differentiation. FGF prevents

premature differentiation of cardiac progenitor cells and regulates heart development during

embryogenesis [208, 209]. Autophagy in cardiac fibroblasts is induced either by serum

deprivation or β2 adrenergic stimulation, which can eliminate misfolded, trimeric forms of

type I pro-collagen produced in error by these cells [210, 211].

Autophagy in Disease Pathogenesis

Autophagy contributes to homeostasis and regulates various physiological processes

by mobilizing intracellular energy sources, degrading entire organelles such as mitochondria,

peroxisomes, ER and by preventing accumulation of altered or misfolded protein that might

otherwise be toxic and might trigger apoptosis [212]. It is the major lysosomal pathway for

the nonselective delivery of cytoplasmic components during periods of starvation or stress

[213, 214]. In certain other conditions autophagy can induce selective digestion of specific

organelles like mitochondria and peroxisomes [215]. There is evidence to support selective

autophagy Atg7-deficient mouse liver displays defects in peroxisome removal, and

chemically-induced endoplasmic reticulum stress leads to endoplasmic reticulum membrane

autophagy disease [216]. Dysregulated autophagy may contribute to the pathogenesis of

diseases [217]. Deficiencies in the autophagic pathway have been linked to several

neurodegenerative diseases such as Huntington’s, Parkinson’s and Alzheimer’s diseases

[218]. Autophagy has also shown to play role in cancer progression [219, 220] and infectious

diseases [221].

In most disease models and histopathological samples of diseased tissue, increased

autophagosome formation has been observed. However, this could be either because of

decreased removal or increased formation of autophagosomes, which relies on the complex

interplay of the various cellular regulators and activators of autophagy [222]. Defects in

42

autophagosome lysosome fusion and genetic or functional alterations causing impaired

delivery of autophagosomes to the lysosome are also implicated in disease pathogenesis. It is

thus possible that impaired autophagic clearance may contribute to the pathogenic

mechanism, for example in dynein mutations causing motor neuron diseases [165].

Accumulation of protein aggregates due to maladaptive autophagy is yet another player

initially correlated in neurodegenerative diseases, but later observed in other diseases as well

[223, 224]. Failure of autophagosomes fusion due to LAMP2 mutations or accumulation of

protein aggregates has also been observed in myodegenerative diseases [222].

Several studies in systemic and tissue-specific Atg mutations in mice have implicated

this protein in maladaptive autophagy causing cancer, and neurodegenerative diseases [225,

226]. Atg5- and Atg7-knockout mice display muscle cell deterioration and progressive

muscle weakness [227, 228]. Atg7-knockout mice also show reduced autophagy, resulting in

pancreatic β-cell degeneration and reduced insulin secretion [229, 230]. Reduced Beclin-1

expression in the brain is correlated with the increased occurrence of aging-related

neurodegeneration [231]. Atg mutations also produce gastrointestinal abnormalities similar to

those found in Crohn’s patients, including Paneth cell dysfunction and impaired granule

secretion [232, 233].

Autophagy in Cardiovascular Diseases

All the above literature signifies the role of autophagy in development, differentiation,

normal physiological processes, and in disease pathogenesis. The heart is critical in

maintaining homeostasis and thus the role of autophagy in cardiovascular pathology takes on

a modicum of importance. In the heart, autophagy serves a predominantly pro-survival

function by removing protein aggregates and damaged organelles under basal condition

[234]. The finding that Atg5 deficiency in murine hearts results in rapid LV dilatation and

contractile dysfunction underlines the important role autophagy plays in maintaining cellular

43

homeostasis in the myocardium [235]. Studies have demonstrated that constitutive autophagy

under basal conditions in the heart is an important mechanism for maintaining cardiomyocyte

size, overall myocardial structure, and proper morphology of mitochondria [236]. The heart

has a high rate of blood flow and contains more mitochondria compared to any other tissue of

the body, which implies a high rate of aerobic respiration for energy generation. Following

myocardial infarction, the heart will have compromised oxygen and nutrient availability due

to ischemia, resulting in upregulation of autophagy to ensure the availability of energy

substrates, recycling amino acids and free fatty acids for energy production [237]. In mice,

caloric restriction is also an inducer of autophagy post-MI, increasing ATP content and

reducing ventricular dilatation [238]. During cardiac ischemia and reperfusion, autophagy can

lead to cell death [239], and is frequently encountered in patients with coronary artery

disease, aortic vascular disease, and congestive heart failure. In models of MI, ischemia

reperfusion injury, and heart failure, adaptive and maladaptive autophagy appears to play an

important role in disease progression [240]. Reperfusion is the next step after long term

ischemia, an attempt to restore the availability of oxygen. However, reperfusion enhances

ROS generation, leading to the opening of the mitochondrial permeability transition pore

(mPTP) and permeabilization of the outer mitochondrial membrane by Bcl-2-like protein 4

(BAX)/Bcl-2-homologous antagonist/killer (BAK) (Bcl-2 pro-apoptotic proteins)[241, 242].

Opening of mPTP also allows an influx of solutes and water, disrupting the mitochondrial

proton gradient and causing depolarization of the mitochondrial membrane. As a result,

mitochondria are unable to generate ATP, and needed to be removed by autophagosomes in a

process known as a mitophagy, which prevents a potentially catastrophic loss of ATP.

Maintenance of ATP levels is one of the important survival mechanisms during ischemia

[241]. To protect from cell death, mitochondrial proteins are ubiquitinated by proteins

phosphatase and tensin homolog (PTEN)-induced putative kinase-1 (PINK-1) and the E3

44

ligase Perkin, allowing their interaction with the cargo protein p62. The complex of p62 and

ubiquitinated mitochondrial proteins is then sequestered by the autophagosome [243, 244].

Although these events prevent immediate cell death, they eventually cause

hypertrophic myocardial growth, increased afterload, and loss of contractile function,

resulting in heart failure. The magnitude of cardiomyocyte hypertrophy is correlated with

activation of autophagic flux. In mice, cardiomyocyte-specific restricted Beclin-1

expressionleads to increased autophagy, increased pathological remodeling, and early

mortality [245], which can be rescued by inducing haploinsufficiency of Beclin-1. Similarly,

HDAC inhibitors attenuate autophagy and blunt cardiomyocyte hypertrophy [246]. Treatment

of cultured neonatal cardiomyocytes with 3MA suppresses autophagy following ischemia

reperfusion and improves cell viability [247]. Similarly, in a rat model of doxorubicin-

induced cardiomyopathy, 3MA treatment is associated with decreased presence of autophagic

vacuoles and significant rescue of cardiac function [248]. The transcription factor GATA4, a

negative regulator of autophagy-related genes, has also been effective in inhibiting

doxorubicin-induced cardiomyopathy [249].

While this recent work indicates that inhibition of autophagy is detrimental in CVD,

and thus supports a maladaptive role of autophagy in disease progression, there are several

other observations where autophagy acts as a protective mechanism. Knockout of Atg5

induces ventricular enlargement and cardiac dysfunction, suggesting autophagy plays a

compensatory mechanism during heart failure. In ischemia, activation of AMPK and its

inhibition of mTOR induce autophagy. Inhibition of autophagy by overexpression of a

dominant negative AMPK, overexpression of Rheb, and overexpression of a dominant

negative form of glycogen synthase kinase 3β, or deletion of one allele of glycogen synthase

kinase-3β results in increased infarct size [250-252].

45

Although animal and cell culture studies provide evidence for both adaptive and

maladaptive roles of autophagy in the heart, most clinical data supports a maladaptive role of

autophagy in disease progression [253]. In two different studies examining explanted hearts

from patients undergoing partial ventriculectomy and transplantation, respectively,

researchers observed numerous autophagic vacuoles within degenerated cardiomyocytes,

signifying autophagy as a prominent source of cardiomyocyte death [254, 255]. Most studies

of autophagy in cardiac diseases have only examined its role in cardiomyocytes. In contrast,

the role of autophagy in fibroblasts, which play a crucial role in heart remodeling, have not

been evaluated much in this context. Recent evidence indicates autophagy may favor fibrosis

by promoting activation of fibroblasts. Our lab has previously demonstrated that increased

production of TGF-β is correlated with increased fibrogenesis and upregulation of autophagy

markers in cardiac myofibroblast in both a rat model of MI as well as human cardiac

myofibroblast derived from scar tissue [256]. Other groups have identified mTORC2

signalingas an inducer of production of the profibrotic growth factor CTGF as well as

fibroblast activation [257]. Aránguiz-Urroz et al. demonstrated that autophagy is involved in

ECM turnover via intracellular degradation of collagen by specific activation of β2

adrenergic receptors [210]. Another study found that increased autophagy correlates with

reduced fibrosis after transaortic constriction, a model of cardiac pressure overload [258].

Aside from its role in cardiomyocytes, there is evidence that autophagy regulates the vascular

smooth muscle cell (VSMC) phenotype, which possesses similar characteristics to

myofibroblasts. Increased autophagy promotes loss of the contractile phenotype and

increased VSMC proliferation, affecting vascular homeostasis [259]. Impaired autophagy

contributes to endothelial dysfunction, and increased production of ROS and inflammatory

cytokines [260]. Studies correlating autophagy in fibroblast and other cell types of

cardiovascular system in disease pathogenesis are rare in the literature and thus the

46

exploration and exploitation of autophagy in this context may provide novel therapeutic

interventions.

Autophagy: A therapeutic target in cardiovascular disease

The recent insights generated in the field of cardiovascular autophagy have raised the

prospect of targeting this cellular pathway for clinical applications. As basal levels of

autophagy help in cell survival and uncontrolled autophagy contributes to pathogenesis, it is

critical to understand the fine balance between these two distinct mechanisms if they are to be

manipulated for therapeutic gain [261]. There are several pharmacological activators and

inhibitors identified as potential regulators of cardiac autophagy [262]. Rapamycin is an

immune suppressive drug known to inhibit mTOR and activate autophagy [263]. Small

molecule enhancers of rapamycin (SMERs),which act independently of the mTOR pathway,

have also been identified [264].5-aminoimidazole-4-carboxamide ribonucleotide (AICAR),

an AMPK analogue which acts upstream of mTOR, stimulates autophagy [265]. Stress

inducers like brefeldin A, thapsigargin and tunicamycin increase the formation of autophagic

vesicles via expression of Beclin-1 and LC-3II, and have been shown to provide

cardioprotection [266]. Drugs targeting mTOR-independent pathways acting on myoinositol

phosphate metabolism have also been evaluated as inducers of autophagy through decreased

inositol triphosphate (IP3) receptor activity, including lithium chloride, sodium valproate,

xestospongin B, and carbamazepine [267, 268]. The FDA has approved two activators of

autophagy, minoxidil (a K+/ATP channel opener) and clonidine (a Gisignaling activator) for

use in humans [269]. Resveratrol is an antioxidant that enhances autophagy by activating

sirtuin, and AMPK is cardioprotective [270]. Metformin, a drug used in diabetes, stimulates

AMPK-dependent autophagy and helps in diabetic cardiomyopathy [271]. Cholesterol

lowering drugs including the various statins may reduce infarct size by activating mitophagy

[272].

47

Amongst the inhibitors of autophagy, 3MA is most well-known. It acts by inhibiting

autophagosome formation through interference of formation of the Vps34/Beclin-1 complex

[148]. It also acts as a PI3K inhibitor, as do two other drugs, wortmannin, and LY294002,

also found to inhibit autophagy [273]. As PI3K signalling is pleotropic in nature and involves

different classes of enzymes, some of which can activate autophagy, the context of its use is

of dire importance in determining its efficacy as a therapeutic. 3MA inhibits the activity of

p38 mitogen-activated protein kinase (MAPK) and c-Jun N-terminal kinase (JNK) as well

[274]. The histone deacetylase inhibitor trichostatin A inhibits autophagy and prevents or

reverses pathological cardiac remodeling by pressure-overload stress.

Suberoylanilidehydroxamic acid (SAHA), a different HDAC inhibitor, activates autophagy

and reduces cardiomyocyte death and infarct size after I/R [246, 275]. Similarly, granulocyte

colony-stimulating factor (G-CSF) treatment in hamsters with cardiomyopathy reverses

excessive activation of autophagy, which correlates with increased survival, improved cardiac

function, and reduced fibrosis [276]. So far, the most specific targets identified in level 3 are

inhibitors of v-ATPase (bafilomycin A1, concanamycinA). Bafilomycin A1 is a specific

inhibitor of v-ATPase in cells, and it inhibits the acidification of lysosomes and endosomes.

As early as 1998, bafilomycin A1 was reported to prevent maturation of autophagic vacuoles

by inhibiting the fusion between autophagosomes and lysosomes [277]. E64d and pepstatin A

are two autophagy inhibitors that function by suppressing lysosomal proteases. E64d is a

membrane-permeable inhibitor of cathepsins B, H, and L, whereas pepstatin A is an inhibitor

of cathepsins D and E.Lysosomotropic agents (able to penetrate the lysosome) such as

chloroquine (CQ) are also a valuable tool for blocking the late stage of autophagy [278]. As a

lysosomal lumen alkalizer (which includes chloroquine, hydroxychloroquine, NH4Cl, and

neutral red), it can also possesses anticancer properties. Lysosomotropic agents are the only

clinically relevant autophagy inhibitors widely used as anti-malarial and anti-rheumatoid

48

agents [279]. A newly developed dimeric form of chloroquine called Lys01 is a highly potent

autophagy inhibitor, ten-fold more potent that of CQ, that contains the spacer N,N-bis-(2-

aminoethyl)-methylamine as a connector between two chloroquine moieties [280]. A newer

water-soluble version called Lys05 is even more potent in its accumulation in the lysosome,

though at high doses has been associated with Paneth cell dysfunction in mice [281].

Sciarretta et al. have proposed several criteria for the selection of dugs targeting

autophagy [282]:

1. Activators or inhibitors of autophagy should be chosen based on the pathological

context.

2. The extent of autophagy modulation should be considered.

3. Autophagy is activated through different signaling mechanisms in different cardiac

diseases which should be understood.

4. Pharmacological modulators of autophagy may exert autophagy-independent functions,

which should be considered.

Targeting autophagy promises to be great therapeutic strategy to treat or prevent heart

diseases. To overcome the drawbacks of pharmacological modulators of autophagy, it is

necessary to explore all the underlying mechanisms in the contexts of CVD and the related

cellular processes involved in progressive diseases such as cardiac fibrosis.

Autophagy in activation of fibroblasts

Through secreting ECM components as well as autocrine and paracrine signaling

molecules that drive and maintain fibrosis, cardiomyocyte hypertrophy, and inflammation,

activated myofibroblasts are key drivers of cardiac fibrosis and cell death [283]. Recently, a

correlation has been established between dysregulation of autophagy and the development of

fibrosis in various organ systems, in the contexts of both up- and down-regulation of

49

autophagic processes [167]. In the liver, hepatic stellate cells were prevented from

differentiating into myofibroblast-like cells due to fibrosis-associated reduction in autophagy

[284, 285]. Autophagy was upregulated during the activation of quiescent hepatic stellate

cells (HSCs), leading to liberation of free fatty acids that undergo mitochondrial β oxidation.

In this mechanism, autophagy handles production of ATP to maintain the activated HSC

phenotype, which like myofibroblasts are hypersynthetic for ECM components and highly

proliferative, leading to hepatic fibrosis [285]. Fibrogenic kidney and lung cells have

demonstrated a reliance on autophagy in maintaining activated and fibrogenic phenotypes. In

a human embryonic fibroblast cell line, autophagy plays a critical role in activation to

fibroblasts[257]. In these cells, prolonged autophagy was associated with inhibition of

mTORC1 activation, and unanticipated augmentation of mTORC2 activation leading to

CTGF-dependent differentiation [257].The finding that inhibition of autophagy via PI3K with

LY294002, wortmannin, or 3MA prevented differentiation, as did inhibition through Atg7

silencing, indicates the critical role of autophagy in the fibroblast activation process.

However, the necessity of autophagy in activation of primary cardiac fibroblasts has not yet

been examined.

50

Rationale and Hypothesis

Rationale

The activation of cardiac fibroblasts to myofibroblasts is the hallmark event for

initiating cardiac fibrosis. As autophagy have been shown to participate in a number of

cardiac pathologies, including MI and is implicated in the induction of fibrogenesis, we

hypothesized (i) that autophagy is the causal for mechanical stress-induced activation of

cardiac fibroblasts to myofibroblasts in vitro and in vivo and (ii) that inhibition of autophagy

will abrogate the fibroblast activation in vitro, and (iii) that inhibition of autophagy in a rat

model of chronic MI will reduce the adverse cardiac remodeling of cardiac ECM in vivo.

51

Hypothesis

General hypothesis: Autophagy is upregulated in heart failure, which causes

fibroblast activation to myofibroblast (Figure 5).

Figure 5: General hypothesis. Panel A. Post-MI cardiac tissue undergoes sequence of

pathological events causing activation of cardiac fibroblast, matrix remodeling and scar

formation. Panel B. Mechanical stress is mimicked in vitro, by plating fibroblast on plastic

tissue culture plates. We hypothesize, that mechanical stress induces enhanced autophagy in

the cardiac fibroblast in vivo and in vitro. Activated autophagy is a positive regulator for

activation of cardiac fibroblasts to myofibroblasts.

A B

52

Specific Hypothesis 1. Mechanical stress induces autophagy and activation of

cardiac fibroblasts in vitro

It has been established that mechanical stress both in vitro and in vivo induces

activation of cardiac fibroblasts to myofibroblasts [286], though this effect has not yet been

studied in freshly isolated P0 fibroblasts. Additionally, mechanical stress has been shown to

induce autophagy in cardiomyocytes [287], and is associated with the activation of

fibroblasts[257]. We hypothesized that mechanical stress concomitantly induces fibroblast

activation and induces autophagy in cardiac fibroblasts in vitro.

Objective 1.1: Determine the effect of mechanical stress on cardiac fibroblasts

Primary unpassaged (P0) primary rat cardiac fibroblasts were subjected to mechanical

stress by plating on non-compressible plastic tissue culture dishes. Time-dependent induction

of autophagy was examined by measuring levels of LC-3I and lipidated LC-3II at 24, 48, and

72 hours post-plating. Activation of cardiac fibroblasts to myofibroblasts was determined by

measuring protein expression of the myofibroblast marker α-SMA at 24, 48, and 72 hours.

Specific Hypothesis 2. Autophagy is necessary for mechanical stress-induced

activation of cardiac fibroblasts in vitro

Inhibition of autophagy has been shown to prevent myofibroblast conversion in

embryonic lung tissue fibroblasts [257], and TGF-β induced conversion of atrial fibroblasts

[256]. We hypothesized that inhibition of autophagic flux with the inhibitors Baf-A1, CQ,

and 3MA would prevent conversion of primary rat cardiac fibroblasts to myofibroblasts.

Objective 2.1:Determine the Effect of Inhibition of Autophagy in Cardiac Fibroblasts

To investigate the correlation of fibroblast activation of primary unpassaged P0 rat

cardiac fibroblasts and autophagy, we subjected these cells to mechanical stress by plating on

non-compressible substrate and treatment with inhibitors of autophagic flux Baf-A1, CQ, and

53

3MA at various concentrations. We then examined autophagic induction by measuring levels

of LC-3I, lipidated LC-3II, and the lysosomal cargo protein p62. Visualization of autophagy

inhibition in cardiac fibroblasts was achieved by immunofluorescence staining for LC3II, and

the presence and accumulation of autophagosomes, determined via transmission electron

microscopy (TEM).

Objective 2.2: Determine the effect of inhibition of autophagy on myofibroblast

phenotype

To determine the effect of autophagy inhibition on development of the myofibroblast

phenotype, we treated mechanically stressed fibroblasts with autophagy inhibitors for 48

hours and examined protein levels of key myofibroblast markers α-SMA and ED-A

Fibronectin, phosphorylation of p38 MAPK, and development of α-SMA stress fibers via

immunofluorescence.


function

To determine the effect of autophagy inhibition on developing myofibroblast function

in mechanically stressed cardiac fibroblasts, we treated cells with autophagy inhibitors for 48

hours and assayed cells for contractile ability (via collagen gel contraction assay) and

migratory ability (via scratch assay) at varying time-points.


function

As MAPK signaling has been shown to play an important role in fibroblast activation,

we determined the effect of mechanical stress on p38 MAPK phosphorylation levels at 24,

48, and 72 hours in the presence and absence of autophagy inhibitors.

54

Specific Hypothesis 3. Inhibition of autophagy in vivo will attenuate cardiac

fibrosis following myocardial infarction

There are numerous lines of evidence indicating that autophagy plays a role in cardiac

remodeling in the subacute stage post-injury in various animal models [288-290], though

studies are lacking in the later stages of cardiac remodeling. Thus, we hypothesized that

inhibition of autophagy in vivo will attenuate cardiac remodeling in post-MI rat hearts after 4

weeks.

Objective 3.1: Determine the effect of inhibition of autophagy on cardiac remodeling

To determine the effect of autophagy inhibition on cardiac remodeling, we utilized a

rat model of MI that employs permanent coronary ligation of the left anterior descending

(LAD) coronary artery and sham-operated control animals. Both groups were further

subdivided into those receiving daily treatment with CQ autophagy inhibitor and those

receiving control injections. We assessed post-MI function using serial echocardiography and

assessed remodeling via biochemical serum analysis and Masson’s trichrome staining for

collagen at 4 and 8 weeks post-MI.

55

Materials and Methods

Animal Ethics

Experimental protocols involving animals were reviewed and approved by the

University of Manitoba’s Animal Care Committee following the Canadian Council of Animal

Care Standards.

Cell Isolation and Culture

Cardiac fibroblasts were isolated by using retrograde Langendorff collagenase

perfusion, as previously described [291]. Juvenile male Sprague Dawley rats (150 to 200

grams) were anesthetized with a ketamine:xylazine cocktail (100 mg/kg ketamine; 10 mg/kg

xylazine) via intraperitoneal injection. Upon loss of limb reflexes, heparin (6 mg/kg) was

administered by intravenous injection in the femoral vein to prevent blood clotting. Hearts

were then excised and placed in a 100 mm cell culture dish containing 10 mL of Dulbecco’s

Modified Eagle’s Medium: Nutrient Mixture F12 (DMEM-F12; Gibco, USA) supplemented

with100 U/mL penicillin, 100 U/mL streptomycin and 1.0 μM ascorbic acid. The hearts were

cannulated to the apparatus from the aorta, and subjected to a 5-minute perfusion of DMEM-

F12 followed by a 6-minute perfusion with Minimum Essential Media Spinner’s

Modification (S-MEM; Gibco, USA) to halt myocardial contraction and promote cell

dissociation. Following S-MEM treatment, hearts were then perfused with DMEM-F12

supplemented with 0.1% (w/v) collagenase type II (Worthington Biochemical Corp., USA)

for 20 minutes to digest the tissue. The digested ventricles were then collected in a 100-mm

culture dish and digested via mincing and incubating in diluted collagenase (0.05% w/v) at

37°C for 10 min. The resulting tissue suspension was then neutralized by adding complete

growth media (DMEM-F12 supplemented with 10% fetal bovine serum/FBS, 100 U/mL

penicillin, 100 U/mL streptomycin, 1.0 μM ascorbic acid; 10% FBS DMEM-F12). The cells

were then filtered through a 40μm cell strainer (ThermoFisher Scientific, USA) to discard

56

undigested tissue; the suspension was collected in 50 mL conical tube and centrifuged at 750

x g for 7 minutes. After centrifugation, the cell pellets were re-suspended in 10 mL of

complete growth media. Lastly, each cell pellet was diluted to a final volume of 30 mL, and

10 mL aliquots were plated in 100mm dishes then incubated for 2 hours at 5% CO2, 37˚Cto

allow for cell adherence. Cell cultures were then washed twice with 1X phosphate-buffered

saline (PBS) and incubated with fresh 10% FBS DMEM-F12 media at 37˚C with 5% CO2

overnight. The next day, cultures were rinsed once with 1X PBS and fresh complete growth

media was added. Once the cultures reached 50-60% confluency they were used for

experimental assays. Unpassaged primary cells are henceforth referred to as P0 cardiac

fibroblasts.

Treatment with Autophagy Inhibitors

Three individual pharmacological inhibitors of autophagy were used in the study:

Bafilomycin A1 (Baf-A1), 3-Methyladenine (3MA), and Chloroquine (CQ) (all from Sigma,

USA).

Optimization of Autophagy Inhibitor Treatment by MTT Assay

The 3-(4, 5-Dimethylthiazol-2-yl)-2,5-Diphenyltetrazolium Bromide (MTT) assay

was used to determine the working concentrations of Baf-A1 and CQ and 3MA.

Approximately 5,000 isolated P0 cardiac fibroblasts were plated in 10% FBS DMEM-F12in

each well of a 96-well plate. For each autophagy inhibitor, a range of concentrations was

applied to a set of wells: 0.25 nM -1.0 nM for Baf-A1, 1.0 mM – 5.0 mM for 3MA, and 25

µM -100 µM for CQ. The assay was performed in triplicates for 24, 48 and 72 hours. After

each time point, 20 µL of MTT solution (5 mg/mL of MTT in PBS, filtered with 0.45 μM

filter) was added to each well and incubated at 37˚C for 3 hours. Further media was removed

and 200 µL of dimethyl sulfoxide (DMSO) was added and mixed via trituration. After

5minutes incubation at room temperature, relative cell proliferation was calculated by

57

measuring optical absorbance at a wavelength of 570 nm using a spectrophotometric plate

reader (Bio-Rad Laboratories, USA). The working concentrations for each autophagy

inhibitor are listed in Table 2.

Table 3: Optimized dosages for autophagy inhibitors. Dosages were determined via MTT

Assay in freshly isolated P0 primary rat cardiac fibroblasts.

Drug Name Optimized Working Concentrations

Bafilomycin-A1 0.5 nM, 0.75 nM

3-Methyladenine 1.0mM, 2.5 mM, 5.0 mM

Chloroquine 25 μM, 35 μM, 50 μM

Protein Isolation from Adherent Cell Cultures

Cells were treated with optimized doses of each drug for 48(for CQ and Baf-A1) or

72 (for 3-MA) hours. After incubation, the cells were rinsed once with 1X PBS, and

mechanically scraped on ice in 1.5 mL of cold PBS. The suspension was collected into 1.5

mL microcentrifuge tubes and centrifuged at 14000 xg for 5 minutes at 4oC. Pellets were re-

suspended in radioimmunoprecipitation assay buffer (RIPA), comprising 1% NP-40, 1.0

mMethylene glycol-bis (β-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA),

supplemented with phosphatase inhibitors (10 mMNaF, 1.0 mMNa3VO4) and protease

inhibitors (P8430 Protease Inhibitor Cocktail; Sigma, USA). Lysates were incubated on ice

for 1 hour, sonicated three times for 10 seconds each and centrifuged at 14,000 x g at 4oC for

15 minutes. The supernatant containing the isolated protein was transferred to fresh

microcentrifuge tubes.

Protein quantification was performed using the bicinchoninic acid method

[292].Samples were diluted 1:4 in RIPA buffer; then, 10 μL of each sample was loaded in

58

triplicate onto a 96-well plate. A set of solutions containing varying concentrations of bovine

serum albumin (BSA; Invitrogen, USA) standards was used to generate a standard curve. 200

μL of a solution of bicinchoninic acid and copper II sulphate (50:1 dilution v/v;

ThermoFisher Scientific, USA) was then added to each well and the plate was incubated at

37°C for 30 minutes. Protein concentrations were determined by measuring the absorbance of

the samples at 560 nm on an iMark microplate reader (Bio-Rad Laboratories, USA).

Western Blot Analysis

Protein samples underwent electrophoretic separation under reducing conditions using

one-dimensional 6-12% (non-gradient) sodium-dodecyl sulphate polyacrylamide gel

electrophoresis (SDS-PAGE). Lanes were loaded with 10-20 μg of protein, which were then

subjected to 150 V of electrical potential for 1.5 hours. Separated proteins were transferred to

polyvinylidene difluoride (PVDF) membranes at 300 mA for 75 minutes at 4°C. Membranes

were blocked in phosphate-buffered saline (PBS) with 0.2% Tween-20 (PBS-T) containing

10% (w/v) skim milk for 1.5 hours at room temperature. Primary antibodies were diluted in

either 3% skim milk in PBS or 5% bovine serum albumin (BSA) in Tris-buffered saline with

0.1% Tween-20 (TBS-T), as per the manufacturer’s instructions. The following primary

antibodies were used: α-SMA (1:5,000; Abcam, USA), ED-A FN (1:1,000; Millipore,

Canada), LC-3β (1:1,000; Sigma, USA), β-tubulin (1:5,000; Abcam, USA), eEF2 (1:1,000;

Cell Signaling, USA), p62 (1:1,000; Cell Signaling, USA), total-p38(1:,1000; Cell Signaling,

USA), and phospho-p38 (1:1,000; Cell Signaling, USA). Membranes were incubated with the

primary antibodies overnight at 4oC, with constant shaking. The following day, membranes

were washed three times with PBS-T, for 10 min each and then incubated with horseradish

peroxidase (HRP)-conjugated goat anti-mouse or anti-rabbit secondary antibodies (1:10,000;

Jackson Laboratories, USA) in PBS-T with 3% skim milk for 1.5 hours at room temperature,

with shaking. After 3-4 washes with PBS-T, protein bands were visualized using ECL or

59

ECL Pico (ThermoFisher Scientific, USA) per the manufacturer's instructions, and developed

on CL-Exposure X-ray film (Life Technologies, USA). Equal protein loading was confirmed

by probing for β-tubulin or eEF2. Films were scanned and digitized using a GS-8000

Densitometer (Bio-Rad Laboratories, USA) and quantified using Quantity One™software

(Bio-Rad Laboratories, USA).

Fluorescence Immunocytochemistry

Primary rat P0 cardiac fibroblasts were seeded onto 6-well tissue culture plates (~2.0

x 105 cells/well) containing glass cover slips and allowed to adhere overnight and then treated

with Baf-A1, CQ, or3MA for 48 hours. Cells were fixed with 4% (w/v) paraformaldehyde

(PFA) in PBS (pH 7.4) for 15 minutes at room temperature, followed by 15 minutes of

permeabilization with 0.1% (v/v) Triton-X 100 in PBS (Sigma, USA). Cells were washed

three times with 1X PBS and blocked for 90 minutes with 10% BSA at room temperature.

Cells were washed once with PBS and incubated overnight at 4oC with α-SMA and LC-3β

primary antibodies (1:500 in 1%(w/v) BSA in PBS). After washing 3 times for 5 minutes

each, cells were incubated with a fluorescent-tagged secondary antibody (Alexa Fluor® 555

anti-mouse or 488 anti-rabbit; Invitrogen; 1:700 in 1%(w/v) BSA in PBS for 90 minutes at

room temperature. Cover slips were mounted onto glass slides using Prolong® Gold antifade

reagent with 4’,6-Diamidino-2-phenylindole (DAPI; Molecular Probes, Life Technologies,

USA). Images were acquired using a Zeiss Axiovert 200M epifluorescence microscope and

analysed with AxioVision software (Carl Zeiss Canada, Canada).

Scratch Assay

P0 rat cardiac fibroblasts were used for cell scratch/wound healing assays. Silicone

inserts (Ibidi, USA) were placed in a 6-well culture dish. A cell suspension of 2 x 105 cells /

70 μL was prepared and added into each chamber of the inserts. The cells were allowed to

adhere and grow overnight in complete growth medium. The following day, the medium was

60

changed to low serum (2% FBS) for 24 hours and then treated with 3MA (5Mm), Baf-A1

(0.75 nM) and CQ (35 µM) for 48 hours. After 48 hours, inserts were removed and imaged

by light microscopy every 6hours until the control cells (untreated) reached 100%

confluency. Data was analyzed by comparing differences between wound width at 0, 12 and

24hours in control and treated groups using Wimasis Image Analysis software (Wimasis

Image Analysis, Germany).

Gel Contraction Assay

Collagen gels were prepared by mixing 7.0 mL of a cold collagen solution

(StemcellTechnologies, Canada) with 2.0 mL of 5X concentrated DMEM-F12 containing 100

U/mL penicillin, 100 U/mL streptomycin and 1.0 µM ascorbic acid(pH 7.4). The final

volume was adjusted to 10 mL using double distilled water (ddH2O). 600 µL of the solution

was added to each well of a 24-well plate and allowed to solidify overnight at 370C in a 5%

CO2 incubator. Approximately 2.0 x 104freshly-isolated P0 rat cardiac fibroblasts were plated

in each well with 10% FBS DMEM-F12 and were allowed to adhere for 2 hours. The cells

were rinsed twice with PBS followed by the addition of 10% FBS DMEM-F12. Cells were

allowed to grow for 24 hours in 10% FBS DMEM-F12, followed by 24 hours incubation in

low serum media (2% FBS DMEM-F12). Following serum deprivation, gels were released

using a circular cutting tool. Subsequently, the cells were divided into four groups: (i)

untreated control group, (ii) TGF-β-treated (10 ng/mL) positive control group, (iii) autophagy

inhibition withCQ (10, 25 and 35 μM), Baf-A1 (7.5 nM), or 3MA(2.5Mm), and (iv)

autophagy inhibition with TGF, CQ (10, 25 and 35 μM) + TGF-β1 (10 ng/mL), Baf-A1 (7.5

nM) + TGF-β1(10ng/mL), and 3-MA (2.5Mm)+ TGF-β1(10ng/mL) treatment groups. Images

were taken immediately post treatment (t=0 hours) as well as at 24 and 48 hours post-

treatment. Gels were analyzed using IDL Measure Gel software (University of Calgary,

Canada) to determine total gel surface area at each time point.

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Transmission Electron Microscopy (TEM)

Cells were fixed in Karnovsky’s Fixative and the pellet was resuspended in 5%

sucrose 0.1M Sorensen’s buffer overnight at 4°C. Cells were pelleted and post-fixed with 1%

osmium tetroxide in 0.1 M Sorensen’s Phosphate Buffer for 2 hours, and dehydrated and

embedded in Embed 812 for sectioning. Semi-thin sections (1 μM) were cut from the blocks

and stained with toluidine blue for inspection, followed by sections of 200 nM which were

placed on copper grids for staining with uranyl acetate and counter staining with lead nitrate.

Imaging was done using a Philips CM10 electron microscope.

Experimental model of myocardial infarction

Adult male Sprague-Dawley rats of 250 – 300 grams body weight were used to create

an experimental model of myocardial infarction (MI). Our lab has extensive previous

experience with this technique[293].Animals were initially divided into two groups,

including(i) coronary ligated animals - animals underwent permanent surgical occlusion of

the left anterior descending coronary artery (LAD), and (ii) a sham-operated group wherein

animals underwent surgery, and the suture was introduced but was not tied, and thus we did

not occlude the LAD. Animals were then subdivided into timed groups, (i) 4-week and (ii) 8-

week post-MI, based on the time-point when the animals were to be sacrificed. To determine

the effect of autophagy inhibitory drug CQ in MI-induced animals, animals underwent

treatment with CQ (40mg/kg) for 3 weeks starting after week 1 post-surgery in the 4-week

model, and treatment for 2 weeks starting from week 6 in the 8-week model. There were 4

groups for each time-point; (i) sham; (ii) sham + CQ treatment (iii) MI, and (iv) MI+CQ

treatment, as is shown in Figure 6. Upon sacrifice, tissues were collected from distinct zones:

viable left ventricle (LV - border zone of infarct) and LV scar (infarct scar) from the ligated

(MI-induced) groups. Complete left ventricles were collected from the sham-operated groups.

Tissues for protein isolation were flash-frozen in liquid nitrogen and stored at-80oC until

62

further processing for various assays. Tissues required for histology and immunofluorescence

were frozen in Optimal Cutting Temperature (OCT) compound blocks and stored at -

80oC.Blood was also collected before and after CQ treatment, and serum was separated for

evaluation of toxicological parameters.

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Figure 6: In vivo Study Timeline. Chloroquine (CQ)-treated left anterior descending (LAD)

ligation rat myocardial infarction (MI) model timeline and treatment groups. A. Treatment

Groups: Adult male rats were randomly divided in two broad groups, sham-operated and

LAD ligated(MI model). Each of these groups was further divided into subgroups of control

(no drug) and CQ-treated. B. Timeline of the study. Rats underwent LAD surgery and after

the first week, both sham-operated and LAD animals received 40mg/kg chloroquine daily for

3 weeks. 4 weeks post-MI these animals were sacrificed and their serum was collected for

biochemical assays. Another group of CQ-treated animals received 40mg/kg CQ daily from 6

weeks onwards until8 weeks and were then subjected to serum collection and sacrifice at the

end of 8 week. Serial echocardiography was performed for 4- and 8-week group animals at

the end of 2,3 and 4 weeks, as well as 6,7 and 8 weeks, respectively.

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Echocardiography

Non-invasive transthoracic echocardiography (TTE) was performed serially on rats

post-surgery at 2, 3, and 4 weeks post-surgery in the 4-week MI-model and 5, 6,7, and 8

weeks post-surgery in the 8-week MI-model. To perform TTE, chest hairs of all the animals

were shaved with an electric razor after anesthesia with 3.0% (v/v) isoflurane gas. The

animals were maintained in this state with 2.0% isoflurane and TTE was performed using an

acoustic gel and 5.0 MHz probe (Vivid 7, GE Medical Systems, USA) in the parasternal short

axis view at the level of the papillary muscles to acquire M-mode images. EchoPAC PC

software (v.112, GE Medical Systems, USA) was used to analyze M-mode images.

Pathological Analysis of Blood Serum

Blood serum was collected from animals subject to LAD ligation or the sham

operation, immediately prior to sacrifice at 4 and 8 weeks, in order to determine effect of CQ

on liver, kidney, and cardiac tissues. Aspartate transaminase/alanine transaminase

(AST/ALT), creatinine, and lactate dehydrogenase (LDH) levels were tested for each organ,

respectively. 1.5 – 2mL of blood was collected in serum separator test tubes. Tubes

containing blood were centrifuged at 2,000 rpm for 15 minutes within one hour of blood

collection. The separated serum was then collected in individual 1.5 mL microcentrifuge

tubes for each test. The serum samples were sent to the pathology laboratory of St. Boniface

General Hospital (Winnipeg, MB, Canada) for analysis. LDH levels were assayed with 2

hours of sample collection, followed by the AST/ALT and creatinine determinations.

Protein Isolation from Whole Tissue

Cardiac tissues isolated from all four treatment group of animals, weighing 30-50mg

were stored at -80oC were used for protein isolation. Tissues were crushed using a motor and

pestle in liquid nitrogen, collected in centrifuge tubes, and mixed into SDS lysis buffer (10μL

for every 1.0 mg of tissue) containing 250 mmol/L Tris-HCl (pH 6.8), 4% SDS, 20% (v/v)

65

glycerol and1.0 mM ethylene glycol-bis (β-aminoethyl ether-N,N,N’,N’-tetraacetic) acid

(EGTA), plus phosphatase inhibitors (10 mMNaF, 1.0 mMNa3VO4) and protease inhibitors

(P8430 Protease Inhibitor Cocktail; Sigma, USA), followed by incubation on ice for one

hour. Samples were then sonicated three times, for 10 seconds each. Lysates were transferred

to QIAshredder columns (QIAgen, Germany) and centrifuged at 14,000xg at 4oC for 2

minutes. The supernatant collected in the bottom of the columns was used for protein content

determination (Smith’s assay, as described above) and Western blot analysis.

Tissue Sectioning for Histology and Immunofluorescence

Cardiac tissue excised from experimental MI-model rats at 4-weeks and 8-weeks

post-MI was stored at -80oCin OCT blocks. Tissue sections of were cut at a 5 μm thickness

using a microtome (Microm HM550 Cryostat; ThermoFisher Scientific). Sections were fixed

with 4% (w/v) PFA in PBS and used for immunofluorescence following the same protocol

used for fixed cells.

Masson’s Trichrome Staining

To determine the distribution of fibrous collagens within the myocardium from in vivo

preparations of hearts, we used Masson’s trichrome staining technique. Cardiac tissue

sections were obtained from frozen OCT blocks and mounted on glass slides. The sections

were blocked in Bouin’s fixative (5% (v/v) acetic acid, 9% (v/v) formaldehyde, 0.9% (w/v)

picric acid) at 60oC in a water bath. The sections were then washed 3 times with PBS to

remove the excess fixative and stained with Weigert’s iron hematoxylin working solution

(Hematoxylin, 95%alcohol, 29%ferric chloride and concentrated HCL) for 10 minutes at

room temperature. The excess solution was rinsed with water for 5 minutes. Next, the slides

were subjected to Biebrich scarlet-acid fuchsin solution 1% beibrich scarlet, 1% fuchsin acid

and glaial acetic acid) for 10-15 minutes. Then sections were washed with water and

subjected to phosphomolybdic-phosphotungstic acid for 10-15 minutes until the red staining

66

for collagens appeared. The slides were then directly stained in aniline blue for 10-15 minutes

and rinsed with water. Next, the slide preps were subjected to1% glacial acetic acid for 5

minutes and washed briefly for 5 minutes. Finally, slide preparations were dehydrated with

95% ethanol for5 minutes Sections were incubated with xylene twice(3 minutes each) and

mounted with coverslips using Permount™ mounting medium (Fisher Chemical, USA).

Images were acquired at 4X using light microscopy.

Hydroxyproline Assay

The Hydroxyproline assay is a semi-quantitative biochemical method for evaluating

tissue collagen content. This assay allows for the detection of hydoxyproline, common to

collagens and elastin, and because collagen content far exceeds elastin in heart tissue, has

been used as a measure of total collagen content in the tissue [294, 295]. The basic principle

of this assay is when oxidized hydroxyproline content in the tissue/serum samples react with

4 dimethylamino benzaldehyde (DMAB), producing a colored product which can be

quantitated by its absorbance at 560nm. We used the Sigma–Aldrich kit according to the

manufacturer’s instructions. We used 10 mg of the LV tissues (control and viable eg, remote

to the infarct scar) in our assays. The tissue was homogenized in 100µl water and subjected to

100 µl of concentrated HCL (12 M) at 120oC for 3 hours. Next the samples were carefully

collected and centrifuged at 13,000xg for 5 minutes in order to receive a clear supernatant.

Further, 5 -10µl of supernatant was added to a 96 well plate and left to evaporate in the oven

at 60 C until they were dry. Next we added 100µl of chloramine oxidation mixture and left

for 5 minutes so that it could oxidize the hydroxyproline. Later added 100 µl of DAMB and

incubated at 60o C for 90 minutes to give colored product. Finally we measured the

absorbance at 560nm and calculated the concentration of hydroxyproline content in our 4

study groups- sham, sham + CQ, MI and MI + CQ.

67

Statistical Analysis

Data are expressed as mean ± SEM. Each n represents a single animal, and all studies

were conducted with at least 3 individual animals in a given group. Statistical significance

between the groups was assessed by Student t-test or where multiple experimental groups are

compared one-way ANOVA followed by Student-Newman-Keuls post-hoc test using

SigmaPlot software (Systat Software, San Jose, CA, USA). A P-value < 0.05 was considered

statistically significant.

68

Chapter 1: Role of Autophagy in Fibroblast Activation In Vitro

Introduction

Autophagosomes have been observed in the myocardium in ischemia/reperfusion

models [17, 18], and at the cellular level, autophagy is involved in epithelial to mesenchymal

transition (EMT) as well as mesenchymal to epithelial transition (MET) [22, 23]. It has been

suggested that the nature of remodeling within cardiac tissue eg, whether it's adaptive or

maladaptive, depends upon the degree of autophagic induction [15]. Both EMT and MET

contribute to differentiation, wound healing and proliferation [23]. The link between cell

differentiation and autophagy has been demonstrated using inhibitors of autophagy.

Treatment with CQ or siRNA selective for ATG3-ATG7 suppresses starvation-induced

autophagy and subsequent EMT through TGF-β/Smad signaling in hepatocarcinoma cells

[22]. Our previous studies of human atrial fibroblasts indicate that the onset of autophagy and

cardiac fibrosis are tightly associated with one another. The use of inhibitors of autophagy

may be useful in revealing the role of autophagic pathways in pathological signaling

processes such as cardiac fibroblast activation to myofibroblasts [13, 16]. When isolated and

cultured in vitro, cardiac fibroblasts undergo rapid activation to myofibroblastic phenotype

which includes among other markers, increased expression of α-SMA [296]. Substrate

stiffness has been found to be a player in this activation [286, 297, 298]. Thus we employ

mechanical stress, in the form of a stiff plating substrate, to induce conversion of cardiac

fibroblasts to the myofibroblast phenotype, as previously published [102, 297, 299].

During autophagy in mammalian cells, autophagosomes engulf components in the

cytoplasm, including proteins and organelles. A soluble microtubule-associated protein

1A/1B-light chain 3 (LC3) is ubiquitously distributed in mammalian tissues as well as

cultured cells, and during engulfment of cytoplasmic contents by autophagosomes, its

cytosolic form (LC-3I) becomes simultaneously conjugated to phosphatidylethanolamine to

69

form LC3-phosphatidylethanolamine (LC-3II). LC-3II is recruited to autophagosomal

membranes, and is degraded alongside autophagosome contents when autophagosomes fuse

with lysosomes to form autolysosomes. Therefore, LC-3II turnover is an indicator of

autophagic activity, and immunodetection of lipidated (phosphatidylethanolamine-

conjugated) LC3 is used as a reliable method for monitoring induction of autophagy [300,

301]. In the final step where the lysosome fuses with autophagosomes, the digestive acidic

enzymes within this compartment degrade the cellular material. In order to successfully

complete autophagy, maintenance of the acidic pH (4.5 to 5) within the autophagolysosome

is very critical and is achieved by the action of membrane bound v-ATPases.

Pharmacological inhibitors such as Baf-A1and CQ interfere with membrane-bound v-ATPase

activity and neutralize the pH within the autophagolysosomes, and thus inhibit autophagy by

preventing autophagosome-lysosome fusion in the final step of the autophagic process [302,

303]. This mechanism is also referred to as autophagic flux inhibition, which results in

accumulation of cellular LC-3II [302]. Conversely, 3MA is a PI3K inhibitor that instead

targets hVPS34 and disrupts the initiation complex, inhibiting autophagy in its initial stages

[304]. We have used these drugs to study the effect of inhibition of autophagy and autophagic

flux in the activation of fibroblasts to hyperactive myofibroblasts. Additional measurement of

autophagic processes, such as levels of the cargo protein p62, which tags and escorts protein

aggregates to autophagosomes, can be measured as further examination of autophagic flux.

Lysosomal degradation of autophagosomes results in decreased p62 levels, and inhibition of

autophagy can induce accumulation of p62 [24].

70

Results

(Part of this work has been published in Oncotarget in October2016. This is an open-access

article distributed under the terms of the Creative Commons Attribution License.)

Title: Inhibition of autophagy inhibits the conversion of cardiac fibroblasts to cardiac

myofibroblasts.

Authors: Shivika S. Gupta, Matthew R. Zeglinski, Sunil G. Rattan, Natalie M. Landry, Saeid

Ghavami, Jeffrey T. Wigle, Thomas Klonisch, Andrew J. Halayko, Ian M.C. Dixon)

Contribution by SSG: Contributed to conceptualization of most of the experiments, carried

out and analyzed all experiments and wrote up 80% of the manuscript)

Induction of Autophagy by Mechanical Stress

Though it has been established that mechanical stress induces autophagy in

cardiomyocytes [287], this effect has not yet been studied in cardiac fibroblasts. Plating on

non-compressible tissue culture dishes induced time-dependent induction of autophagy in

freshly isolated unpassaged (P0) rat cardiac fibroblasts. Activation of cardiac fibroblasts to

myofibroblasts were confirmed by a significant increase in protein levels of α-SMA after 48

and 72 hours of plating, compared to 24 hour controls. Concomitantly, we observed a

significant 6-fold increase in lipidated LC-3II protein levels after as little as 48 hours after

plating, as compared to 24 hour controls. By 72 hours post-plating lipidated LC-3II still

maintained a 5.5-fold significant increase (P< 0.05) compared to the 24 hour control (Figure

7A and B).

71

Figure 7: Temporal activation of autophagy and fibroblast in P0 adult rat cardiac

fibroblasts. Panel A. There was a significant increase in the level of the autophagosome

marker, LC-3II at 48 and 72 hours post-plating on a non-compressible plastic substrate vs. 24

hours. Panel B. Western blot analysis for the myofibroblast marker α-SMA showed a

significant increase 48 and 72 hours after plating when compared to 24 hour controls. Data

are mean ± SEM (n = 3) (*P< 0.05 24 hours vs. 48 hours; #P< 0.05 24 hours vs. 72 hours;

φP< 0.05 48 vs. 72 hours).Significance was determined using one-way ANOVA followed by

a Student-Newman-Keuls post-hoc test.

72

Inhibition of Autophagy in Cardiac Fibroblasts

To determine the relationship between cardiac fibroblast activation and autophagy in

primary, unpassaged P0 rat cardiac fibroblasts, we treated cells with two inhibitors of

autophagic flux, bafilomycin-A1 (BafA1) and chloroquine (CQ), and with 3-methyladenine

(3MA), which inhibits autophagy initiation. Levels of LC-3I and lipidated LC-3II were

assayed via Western blotting, revealing a significant increase in LC-3II lipidation with Baf-

A1 and CQ treatment (P< 0.05) compared to controls (Figure 8A, B and C). No significant

change in lipidated LC-3II levels was observed with 3MA treatment. Autophagy inhibitors

have also been found to induce accumulation of p62 protein levels [24], and indeed, with CQ

treatment, we observed a 3.4-fold significant (P< 0.05) increase in p62 levels at 50 µM CQ as

compared to controls (Figure 8D).

73

#

A

B

C

#

B

A

74

Figure 8: Baf-A1, CQ and 3MA treatment inhibits autophagy in P0 cardiac fibroblasts.

Panel A and B 48 hours of Baf-A1 treatment (5.0 nM and 7.5 nM) showed no significant

change in LC-3I protein expression. However, there was a significant increase in the

expression levels of LC-3II with 7.5 nM Baf-A1 vs. untreated controls. Cells treated with CQ

(25 µM and 50 µM) for 48 hours demonstrated increased accumulation of LC-3II vs.

untreated controls. The 50 µM CQ treatment group was significantly increased vs. control

while the 25 µM group was trending upward but did not reach significance. Panel C.

Inhibition of autophagy in the 3MA treatment group showed no significant change in LC-3II

or LC-3I. Panel D. p62 protein levels were significantly increased following CQ (50 µM)

treatment vs. untreated control. Data are mean ± SEM (n=3-4) (#P< 0.05 control vs. 7.5 nM

Baf-A1 and 50 µM CQ). Significance was determined using one-way ANOVA followed by a

Student-Newman-Keuls post-hoc test.

D

75

We utilized transmission electron microscopy (TEM) to visualize autophagosomes in

cardiac fibroblasts treated with autophagic flux inhibition drugs (Baf-A1 and CQ).This

confirmed autophagosome presence and accumulation in both treatment groups compared to

controls (Figure 9A and B).

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Figure 9: Ultrastructure of rat ventricular cardiac fibroblasts. TEM confirms the

presence of accumulated autophagosomes as compared to time matched controls in those

cells treated with Baf-A1 (5.0 nM and 7.5 nM; Panel A) and CQ (25 µM and 50 µM; Panel

B). The magnification for all images are 5000 and 10,500, 19,000 or 30,000. The scale is

A

B

77

shown on each image. Data are mean ± SEM (n=3-4) (#P< 0.05 control vs. 7.5 nM Baf-A1

and 50 µM CQ).

78

Inhibition of Autophagy and Myofibroblast Features

The concomitant increase in myofibroblast marker levels and lipidated LC- II lead us

to question whether inhibition of autophagy may affect the process of activation in P0 cardiac

fibroblasts. Using Western blot analysis, we determined that all three autophagic inhibitors

significantly reduced (P< 0.05) protein levels of myofibroblast markers α-SMA and ED-A

FN (Figure 10A, B and C). Treatment with 50 µM CQ completely abolished expression of α-

SMA and ED-A FN.

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Figure 10: Inhibition of autophagy reduces expression of key cardiac myofibroblast

markers. Western blot analysis of P0 cardiac fibroblasts treated with autophagy inhibitors

show decreased expression levels for ED-A FN and α-SMA. Panel A. 48 hours of Baf-A1

(5.0 nM and 7.5 nM) treatment resulted in a significant reduction in α-SMA (40 % and 70%)

and ED-A FN (35% and 50%) levels respectively. Panel B. 50 µM CQ treatment for 48

hours shows a significant decrease in α-SMA and in ED-A FN levels to near undetectable

levels. Panel C. Treatment groups of 1 mM, 2.5 mM and 5 mM 3MA for 72 hours show

decreased expression levels of α-SMA and ED-A FNvs. controls. Data are expressed as mean

± SEM (n=3-4) (*P< 0.05 control vs. 5.0 nM Baf-A1, 2.5 mM 3MA; #P< 0.05 control vs. 7.5

nM Baf-A1, 5 mM 3MA, 50 µM CQ). Significance was determined using one-way ANOVA

followed by a Student-Newman-Keuls post-hoc test.

C

82

Further examination of autophagic flux via immunofluorescence staining for α-SMA

revealed a decrease in α-SMA-positive stress fibre formation (a key feature of the

myofibroblast) following treatment with all three inhibitors, as well as increased punctate

LC-3β staining, compared to controls (Figure 11A, B and C).

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Figure 11: Immunofluorescence staining of P0 cardiac fibroblast. Immunofluorescence

supports the Western blot data and shows decreased α-SMA stress fibre formation in Baf-A1

CQ and 3MA treated cells vs. control. Baf-A1 (A) and CQ (B) show increased LC-3II

accumulation due to inhibition of autophagic flux vs. control. 3MA (C) treated cells exhibit

decreased LC-3II staining vs. untreated control cells. Scale bar = 10µm (A); 20µM (B); 50µm

(C).

C

86

Inhibition of Autophagy and Myofibroblast Function

We sought to further examine the effect of autophagy inhibition on not only

myofibroblast morphology, but also myofibroblast function – namely contractility and

motility, both of which are increased in the wound healing response and subsequent

activation of fibroblasts to myofibroblasts. To assess contractility, we used a collagen gel-

based contraction assay to determine surface contractility of cells as previously described by

our group and others, utilizing TGF-β1 as a potent inducer of contraction [50, 305]. Treatment

with Baf-A1 (7.5 nM) and CQ (25 and 35 µM), but not 3MA, significantly (P< 0.05)

inhibited TGF-β1-induced contraction after 72 hours (Figures12A, B and C), as compared to

TGF-β1 stimulated positive controls.

87

A

B

88

Figure 12: Effect of autophagy inhibition on cellular contractility of P0 cardiac

fibroblasts. Panels A, B and C. Gel contraction assay of P0 cardiac fibroblasts treated with

7.5 nM Baf-A1 (12A), 10-35 μM CQ (12B) and 5mM 3MA (12C) with and without 10 ng/ml

TGF-β1 for 48 hours. Solid black bars represent untreated and white bars represent TGF-β1

treated cells. Both gel images and histographic representation of the data demonstrate that 7.5

nM Baf-A1 and 35 μM CQ significantly inhibited fibroblast mediated gel contraction vs.

untreated controls Baf-A1 + TGF-β1 and CQ + TGF-β1 treatment also shows a significant

decrease in contraction as compared with TGF-β1 treated positive control. There was no

significant difference in 3MA treated gel contraction assay. White arrows show on

contraction with Baf-A1 and CQ treatment, even in the presence of TGF-β1 in the wells. Data

are mean ± SEM (n = 3- 4) (ϕP< 0.05 comparison of TGF-β1 (-) vs. TGF-β1 (+) groups; *P<

0.05 for control vs. 7.5 nM Baf-A1 treatment, and control + TGF-β1vs. 25 μM CQ + TGF-β1;

#P< 0.05 for control + TGF-β1vs. 7.5 nM Baf-A1 + TGF-β1 and control + TGF-β1vs. 35 μM

C

C

89

CQ + TGF-β1). Significance was determined using one-way ANOVA followed by a Student-

Newman-Keuls post-hoc test.

90

We used a traditional scratch assay to determine the relationship between autophagy

and cardiac fibroblast migration. These assays were performed in the presence and absence of

autophagic inhibitors and migration measured via time required for fibroblasts to fill the

empty wounded area (eg, migrate). Compared to time 0 hour controls, by 12 hours there was

reduced fibroblast migration with inhibition of autophagy 3MA, Baf-A1 and CQ; Figure 13).

After 24 hours, CQ continued to inhibitfibroblast migration, compared to untreated controls

which had fully covered the denuded area. With 35 µM CQ treatment, an approximate 3-fold

and 6-fold reductions in cell migration were observed after 12 and 24 hours, respectively.

Although Baf-A1 treatment appeared to reduce cell migration at the 12 hour time-point via

visual observation, there was not a statistically significant difference in migration. Treatment

with 3MA did not significantly alter cardiac fibroblast migration, even after 24 hours (Figure

13).

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Figure 13: Scratch assay to assess cellular migration of cardiac fibroblasts in the

presence and absence of 7.5 nM Baf-A1, 5mM 3MA, 35 μM CQ. The solid black bars

represent untreated controls, grey bars Baf-A1 treated and white bars represent CQ treated.

Migration was gauged by the gradual population with cells migrating into the cell free zone

on the glass slide over time. Migration of fibroblasts in control (untreated cells) and in cells

treated with 5mM 3MA, 7.5 nM Baf-A1 or 35 μM CQ are depicted in the images. 12 hours

after the initiation of the assay we found a significant decrease in migration in CQ treated

cells when compared to untreated controls. Similarly, after 24 hours, CQ treated cells were

found to migrate more slowly than untreated controls. Control groups were ~ 100% confluent

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at 24 hours. (*P< 0.05 for 0 hours vs. 12 hours and 24 hours in untreated controls; σP< 0.05

for 0 hoursvs. 12 hours and 24 hours in 5 mM 3MA treated groups;φP< 0.05 for 0 hours vs.

12 hours and 24 hours in 7.5 nM Baf-A1 treated groups;#P< 0.05 for 0 hours vs. 12 and 24

hours in 35 μM CQ treatment groups; †P< 0.05 for Baf-A1 vs. CQ 35 μM at 24 hours;

ϕP<0.05 for untreated control vs. CQ 35 μM, treated groups within the same time point). Data

are mean ± SEM (n = 3- 4). Significance was determined using one-way ANOVA followed

by a Student-Newman-Keuls post-hoc test.

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Inhibition of Autophagy and Mitogen-Activated Protein Kinase (MAPK) Signaling

MAPK signaling is implicated in the acquisition of the myofibroblast phenotype, eg

the activation of cardiac fibroblasts. Our preliminary study of the effects of MAPK signaling

on myofibroblast phenotype included Western analysis of P42/44 (ERK) and p38 expression

in both basal and phosphorylated forms. When we compared plating duration (24h, 48h and

72h) among groups, we found no changes in ERK phosphorylation. We also used Western

blotting to test for phosphorylation (eg, activation) of p38 MAPK in unpassaged P0 cardiac

fibroblasts over 72 hours (Figure 14). At both 48 and 72 post-plating, we observed a

significant (P< 0.05) reduction of 80-90% in the ratio of phospho-p38: total p38 levels

(Figure 14A) compared to 24 hour controls. Inhibition of autophagic flux with CQ (50 µM)

resulted in a 12-fold increase in phospho-p38: total p38 levels after 48 hours, comparable to

unpassaged P0 fibroblasts 24 hours post-plating (Figure 14B).

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Figure 14: Effect of cell plating and CQ treatment on p38-MAPK phosphorylation in P0

primary cardiac fibroblasts. Panel A. Phosphorylation of p38-MAPK was detected at 24, 48

and 72 hours after plating of P0 cardiac fibroblasts onto a non-compressible plastic substrate.

Total p38-MAPK was also analysed. Phospho-p38 (P-p38) expression was significantly

decreased after 48 and 72 hours of culture when compared to 24 hour controls. Panel B.

Western blot images indicate an increase in phosphorylation of p38-MAPK in the presence of

50 µM CQ treated for 48 hours, as compared to untreated controls. Lane protein loading was

normalized using β-tubulin. Data are mean ± SEM (n=3) (*P< 0.05 applies to the

comparisons of 24 hours plated cells vs. 48 hours plating; #P< 0.05 applies to the comparison

of 24 hours plating vs. the 72 hours plated group; φP< 0.05 for control vs. 50 µM CQ).

Significance was determined using one-way ANOVA followed by a Student-Newman-Keuls

post-hoc test.

B

A

95

To further investigate the role of MAPK signaling in autophagy and induction of

myofibroblast phenotype in cardiac fibroblasts, we utilized a p38 MAPK inhibitor SB203580,

both in the presence and absence of autophagic inhibitors. Pre-treatment with 10 µM

SB203580 show no change in p38 phosphorylation, as compared to control.(Figure 15A).

Combination treatment with both SB203580 + CQ induced a significant increase in p38

phosphorylation, although this increase did not differ significantly from that of CQ alone

(Figure 15A). To examine the effect of MAPK and autophagic inhibition, we examined their

effects on autophagic flux via Western blotting for lipidated LC-3β II levels. Treatment with

SB230580 both in the absence and presence of CQ had no effect on lipidated LC-3β II levels,

though CQ treatment alone showed a significant increase in lipidated LC-3β II (P < 0.05;

Figure15B). Similar effects were observed when SB203580 was used in the presence and

absence of 7.5 nM Baf-A1 (Figure 16A and B).

We then examined the effects of SB203580 in the presence and absence of autophagy

inhibitors on the on fibroblast activation within P0 cardiac fibroblasts. Treatment with

SB203580 alone did not significantly affect protein levels of the myofibroblast markers α-

SMA and ED-A FN compared to untreated controls (Figure 15C and D). Co-treatment with

SB203580 + CQ significantly (P < 0.05) reduced myofibroblast marker protein levels

compared to both untreated and SB203580 alone-treated groups (Figure 15C and D). No

additive effects were observed with SB203580 in conjunction with CQ treatment, as the

addition of SB203580 to CQ inhibition did not result in any significant difference in the

amount of decrease of α-SMA and ED-A FN. Again, similar results were observed with

SB203580 and Baf-A1 co-treatment (Figure 16C and D). Between the two inhibitors of

autophagic flux, CQ had a more pronounced effect on increasing p38 phosphorylation and

repressing the myofibroblast phenotype.

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A

B

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Figure 15: Effect of MAPK P-38 inhibition on primary rat cardiac fibroblasts with and

without CQ treatment. Panel A. Western blot analysis of P0 fibroblast exhibit increased

expression of phospho-p38 levels in CQ and CQ + SB203580 treatment groups when

compared to control and SB203580.Panel B. Treatment of cells with SB203580 revealed

significant increase in expression of LC-3II suggesting increase in autophagy vs. control.

Increased LC3-II expression levels in CQ and CQ + SB203580 treatment groups vs. control,

may reflect accumulation of LC-3II due to inhibition of autophagic flux by CQ. Panels C

and D. Similar to the finding above we found a decrease in phenotypic markers α-SMA and

ED-A FN with CQ and CQ + SB203580 treated cells vs. control and SB203580treated cells.

C

D

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SB203580 treatment was associated with similar expression of both phenotypic markers

when compared to control. Data are expressed as mean ± SEM and representative of 3

individual animals and experiments (eg, n=3) (*P<0.05 control vs.50µM CQ or CQ +

SB203580; #P<0.05 SB203580vs.50µM CQ or CQ + SB203580). Significance was

determined using one-way ANOVA followed by a Student-Newman-Keuls post-hoc test.

99

A

B

C

100

Figure 16: Effect of MAPK inhibition on Baf-A1 treated and untreated P0 cardiac

fibroblasts. Panel A. Phospho-p38 levels show significant increase in Baf-A1 and Baf-A1 +

SB203580 treated cells vs control and SB203580 treated cells. Panel B. Autophagy marker

LC-3II exhibit increased expression in SB203580 treated vs. control group. Baf-A1 treatment

in presence and absence of SB203580 show accumulation of LC-3II suggesting inhibition of

autophagic flux compared to control. Panels C and D. Phenotypic markers α-SMA and ED-

A fibronectin show significant decrease in expression pattern when compared to control and

SB203580. Data are expressed as mean ± SEM and representative of 3 individual animals

and experiments (eg, n=3) (*P<0.05 control vs.7.5nM Baf-A1 or CQ + SB203580; #P<0.05

SB203580vs.7.5 nM Baf-A1 or Baf-A1 + SB203580). Significance was determined using

one-way ANOVA followed by a Student-Newman-Keuls post-hoc test.

D

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Discussion

Wound healing and tissue fibrogenesis are processes requiring the activation and

migration of fibroblasts, as well as the hypersynthetic and contractile abilities of

myofibroblasts. Fibroblast activation to myofibroblast is a critical hallmark of both processes,

of which TGF-β has been shown to be a major driver [306]. TGF-β signaling in fibrosis

occurs both through a canonical Smad-dependent pathway as well as TGF-β - linked p38-

MAPK signaling [307]. Recently, our group demonstrated that autophagy may regulate TGF-

β1-induced fibrogenesis in primary human atrial myofibroblasts [256].

Autophagy in eukaryotes is associated with enhanced longevity [308], as well as

cytoprotection in fibroblasts. Embryonic mouse fibroblasts overexpressing the pro-

autophagic ATG5 gene were found to better resist cellular stress [309]. Additionally,

autophagy is downregulated in fibrotic lesions from various tissues including the lung and

liver [310, 311]. Autophagy has been extensively examined in the context of cardiomyocyte

function and death in response to pathological stimuli, especially in myocardial remodeling.

Cardiomyocyte loss (both apoptotic and non-apoptotic) are crucial in cardiac remodeling and

the development of heart failure, and have been demonstrated in association with oxidative

stress and increased autophagy in ischemia/reperfusion and pressure overload cardiac models

[312, 313]. In both models, inhibition of pro-autophagic Beclin-1 ameliorated adverse cardiac

remodeling. In fact, autophagic inhibition has been shown to alleviate fibroproliferative

events in numerous tissues [314]. However, these studies only describe the autophagic

response before and after disease development, while the current results target the importance

of autophagic flux in the fibroblast activation in freshly isolated, unpassaged P0 rat cardiac

fibroblasts. Studies such as these are rare within the literature due to the technical hurdles

associated with experimentation involving quiescent P0 cells, and thus the need for the

current study is rationalized.

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We utilized an established and highly effective method of inducing fibroblast

activation, that is, plating on hard substrates [4, 14, 27, 33]. Even in primary rat hepatic

stellate cells from TGF-β-null mice, culture on stiff substrates induces the myofibroblast

phenotype [315]. In turn, long-term mechanical strain in skin and heart tissues further

potentiates scarring and fibrosis [297, 316]. Numerous studies have demonstrated rapid

induction of the myofibroblast phenotype in primary fibroblasts plated on non-compressible

tissue culture substrates [296, 305, 317]. Within 16 - 48 hours of plating, our primary

unpassaged P0 rat cardiac fibroblasts exhibited significant increases in protein levels of key

myofibroblast markers α-SMA and ED-A FN, as well as increased presence of α-SMA stress

fibers, an effect that persisted to 72 hours post-plating. The combination of these findings

provides a strong body of evidence supporting the recapitulation of mechanical stress-

induced fibroblast activation in our P0 cardiac fibroblasts. This stiffness-induced fibrobast

activation was accompanied by increased expression of lipidated LC-3β II at both 48 and 72

hours post-plating. This agrees with previously published data from our lab which pointed to

a causal effect of TGF-β1 stimulation in induction of autophagy and cell activation. Cellular

activation is closely linked to increased ECM synthesis in both post-coronary artery bypass

graft (CABG) human atrial and post-MI rat ventricular cardiac tissues [318]. Additionally,

studies in hepatocytes and hepatic carcinoma cell metastasis have demonstrated a

requirement for autophagy in hepatocyte phenotype plasticity and TGF-β1 stimulated EMT

[319].

As autophagy has been described by our lab [256] and others [301] as a necessary

cellular process in fibrotic tissues, we sought to elucidate associated mechanisms in cardiac

fibroblasts activation to myofibroblasts. Western blotting for accumulated p62 and lipidated

LC-3β II levels confirmed successful inhibition of autophagy with all three inhibitors (Baf-

A1, CQ, and 3MA). Via TEM, we confirmed autophagic inhibition through the observation

103

of autophagosome accumulation. A novel finding of this work is the prevention of cardiac

fibroblast activation to myofibroblasts with autophagic inhibition, as demonstrated by

decreased Western blot protein levels of the markers α-SMA and ED-A FN in comparison to

untreated control plated on stiff substrates. Moreover, we determined that CQ was the most

effective of the three drugs in inhibiting activation of cardiac fibroblasts. This is in agreement

with a study by He et al. [320], which showed CQ treatment of hepatic stellate cells in vivo in

a liver fibrosis model was able to diminish α-SMA expression and cellular organization.

However, no studies to date have demonstrated the role of autophagy in repressing cardiac

fibroblast activation, as well as myofibroblast function (specifically cell contraction and

migration), though there is data for other cell types [227, 321, 322]. For example, ATG5

knockout cardiomyocytes exhibit decreased contractile force, and loss of ATG7 in skeletal

muscle induces myocyte loss and reduce force production [227]. Our finding that inhibition

of autophagic flux with either, Baf-A1 and CQ abrogates cardiac myofibroblast contraction

agrees with these findings. In terms of migration, the effects of autophagy have been

examined in HeLa and other transformed cell types [301, 323]. In HeLa cells, ATG3, -5, or -

7 knockdown promoted increased migration compared to competent cells [301]. In MDA-

MB-231 breast cancer cells, autophagic activation with trifluoperazine treatment inhibited

cell migration [323]. These reports contrast with our present findings, in which we find

inhibition of autophagic flux with either Baf-A1 or CQ inhibited migration of cardiac

fibroblasts over 24 hours. One potential reason for this difference in findings may be that

genetic inhibition of ATG gene expression does not specifically target autophagic flux. This

discrepancy may also be the result of vastly different cellular models (our primary,

unpassaged cells vs. immortalized/cancer cell lines). On the other hand, a study in MGC803

gastric cancer cells showed that CQ treatment inhibited cell migration in a dose-dependent

manner [324], which agrees with our findings. It is important to note that CQ possesses non-

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specific inhibitory actions on Toll-like receptor 9 (TLR9), a key factor in cell migration [324]

– thus, although the decreased migratory response to CQ treatment in cardiac fibroblasts was

not anticipated, it may be at least partially attributed to non-specific effects of CQ on TLR9

function. It is interesting to note that 3MA treatment did not significantly affect

myofibroblast functions (contractility and migration). This may be due to the different

inhibitory mode of 3MA, which instead of targeting autophagic flux, blocks formation of

autophagosomes via inhibition of PI3K. Thus, it may be that certain phases of the autophagic

process are more critical for the process of cardiac fibroblasts activation than others.

Nonetheless, inhibition of autophagic flux with either Baf-A1 or CQ prevented not only

morphological phenoconversion features, but also critical myofibroblast functions –

suggesting autophagy is an important process in development of the cardiac myofibroblast.

Concurrent with autophagy induction and activation of the myofibroblast phenotype

(48 hours post-plating), we also observed a significant decrease in phospho-p38: total p38

ratio. Inhibition of autophagy with CQ resulted in an increased expression level of phospho-

p38: total p38 similar to that observed in untreated P0 cardiac fibroblasts at 24 hours after

plating. This data show an inverse relationship between phospho-p38 and markers of both

fibroblast activation and autophagy. Therefore to establish a causal relationship for p38

activation we sought to determine the effect of inhibitors of autophagy in the presence and

absence of an inhibitor of p38 MAPK activity and assess both markers of myobiroblast. To

examine the effect of p38 phosphorylation and autophagic flux on fibroblast activation we

treated cells with SB203580 alone as well as a co-treated CQ or Baf-A1and SB203580for 48

hours. As expected we found that SB203580 treatment alone, which means phospo-p38

inhibition, did not alter the expression of α- SMA and ED-A FN. Next we found co-treatment

of P0 primary fibroblasts resulted in a reduction of the myofibroblast phenotype compared to

controls. We anticipated there should have been a higher degree of expression of α-SMA and

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ED-A FN in the co-treatment groups when compared to CQ and Baf-A1 treatment alone.

However, there was no difference in the level of reduction between CQ or Baf-A1 treated and

CQ or Baf-A1 + SB203580 treated. Which could be due to use of two pharmacological

inhibitors used together. Possibly the Phospho-p38 inhibitor could not work along in

combination with CQ or Baf-A1. Thus to explore the effects of MAPK and autophagy

inhibition simultaneously a genetic KO of p38 along with CQ or Baf-A1 should be tested.

Although phosphorylation of p38 MAPK is clearly related to inhibition of autophagy, the

functional significance of this event remains unknown. In a rat model of hepatic

ischemia/reperfusion injury, Fang et al. [48] also observed a significant increase in p38

phosphorylation with CQ treatment as soon as 6 hours and up to 48 hours after treatment,

which supports our current findings.

A schematic of these findings in primary unpassaged P0 rat cardiac fibroblasts –

plating on stiff substrates, fibroblast activation, the induction of autophagy, and the effect of

inhibition – is shown in Figure 16. It can be concluded that mechanical stress induces both

fibroblast activation and autophagy in cardiac fibroblasts, and inhibition of autophagy blocks

autophagolysosomes formation and results in significant repression of the cardiac

myofibroblast phenotype, findings that support our previous work [305].

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Figure 17: Effect of inhibition of autophagy on primary cardiac fibroblast-to-

myofibroblast phenoconversion. The plating of primary cardiac fibroblasts on non-

compressible plastic matrix is associated with the mechanical induction of autophagy and

decreased phosphorylation of p38, these events attends their activation and phenoconversion

to myofibroblasts. The addition of autophagic flux inhibitory drugs CQ and Baf-A1blocks the

107

fusion of the autophagosome and lysosome, a key component of autophagy, concomitantly

inducing p38 phosphorylation. The mechanism of inducing p38 phosphorylation by CQ and

Baf-A1 is currently unknown. Comprehensively, autophagic flux inhibition significantly

suppresses the myofibroblast phenotype.

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Chapter 2: Autophagy and Cardiac Fibrosis In Vivo

Introduction

Myocardial remodeling is a key event after myocardial infarction (MI). One of the

important hallmarks of cardiac remodeling is the activation of cardiac fibroblasts to

hypersecretory myofibroblasts [101]. Fibroblasts comprise the largest cell population in the

myocardium, arising from various sources, and are principally involved in producing

structural proteins such as fibrillar collagens, which form much of the cardiac extracellular

matrix (ECM) [62]. Cardiac injuries (such as MI) trigger fibroblast activation to

myofibroblasts [119]. After MI, the heart undergoes wound healing wherein fibrillar collagen

deposition contributes to the infarct scar [65]. While this may be viewed as a beneficial event

in the short-term post-MI heart, chronic uninhibited and inappropriate scarring often occurs

post-MI and the cells responsible go on to infiltrate the non-infarcted tissue. Reduced

integrity and elasticity of cardiac tissue results in thickening and stiffening of the remnant or

surviving myocardium, which is also referred to as cardiac fibrosis [325]. Cardiac

myofibroblasts play a key role not only in reparative fibrosis post-MI, but also in

hypertrophy-induced remodeling and fibrosis. Phenoconversion to myofibroblasts is

promoted by growth factors such as transforming growth factor beta (TGF-β), platelet-

derived growth factor (PDGF), and connective tissue growth factor (CTGF), vasoactive

peptides such as angiotensin II (ANGII), as well as a variety of other profibrotic cytokines

released by cardiomyocytes and non-myocyte cells in the injured heart [94]. Physical changes

in the ECM resulting from cardiac injury doubly promote remodeling by allowing the release

and activation of latent TGF-β as well as exerting mechanical tension that has also been

shown to promote fibroblast activation [326, 327]. This activation of fibroblasts shifts the

balance to favour ECM synthesis, leading to ECM thickening and contraction, increasing

synthesis and accumulation of fibrotic depositions that can replace the myocytes and/or

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interrupt the myocyte–myocyte interactions in the myocardium [328, 329]. Enhanced

stiffness of the myocardium impedes ventricular contraction and relaxation as well as

electrical signaling between cardiomyocyte, leading to distorted architecture, impaired

systolic and/or diastolic function of the heart, and arrhythmias [330].

Recent evidence from our lab suggests that autophagy favours fibrosis by enhancing

activation of cardiac fibroblasts to myofibroblasts [256]. Autophagy is associated with

acquisition of markers of fibroblast activation including increased protein levels of

ACTA2/α-SMA (actin-α-2/α-smooth muscle actin), enhanced expression of collagens, and

the formation of stress fibres, which impart contractility for proper wound contracture [256,

288]. This work demonstrated that autophagic fibroblasts also have increased expression and

secretion of CTGF. Inhibition of autophagy using mTOR inhibitor, class I phosphoinositide

3-kinase and class III phosphatidylinositol 3-kinase (PI3K) inhibitors, or through ATG7

silencing, prevented fibroblast activation and attenuated LV remodeling in a mouse model of

MI. In spite of these findings, the mediators linking autophagy to fibroblast activation and

fibrosis still remains largely undefined.

In Chapter One, we demonstrated a link between autophagy and fibroblast activation

using an in vitro mechanical stress model of rat cardiac fibroblast activation (eg, plating on a

noncompressible matrix). One of the critical components for autophagy is the proper

functioning of the lysosome and/or promoting its fusion with autophagosomes. Thus, we

hypothesized that the autophagic insult observed during MI could be attenuated using

chloroquine, which is a lysosomal lumen alkalizer that blocks the autophagy progress by

impairing lysosome formation. Chloroquine and its analog hydroxychloroquine are the only

clinically relevant autophagy inhibitors that are widely used as anti-cancer, anti-malarial and

anti-rheumatoid agents [331]. Recently, Ohm et al. (2014) demonstrated that chloroquine can

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inhibit TLR9-mediated inflammatory responses in murine cardiac fibroblasts. Of interest,

altered TLR 9 signaling has pathophysiological significance both in various cardiovascular

disorders and cardiac remodeling [332].

As we established the efficacy of chloroquine as an anti-fibrotic agent in our in vitro

model of cardiac fibroblast activation to myofibroblasts, we sought to determine the effects of

chloroquine at the preclinical level using in vivo rat model of MI induced via permanent

ligation of the left anterior descending artery (LAD). Interruption of blood flow via LAD

ligation leads to ischemia, mimicking MI and leads to the development of a fibrotic scar

tissue. This surgical procedure imitates the pathobiological and pathophysiological aspects

occurring in MI and thus is apt for the study of protective effect of chloroquine in post-MI

cardiac remodeling.

Methodology

As described in the Materials and Methods section above, we utilized permanent

surgical occlusion of the LAD in rats as an experimental model of MI, and sham-operated

controls underwent surgery without permanent occlusion of the LAD. Timed myocardial

infarction (MI) and sham treatment groups received 40 mg/kg/day injected chloroquine

(CQ) either for 3 weeks (from week 1 in the 4-week model) or 2 weeks (from week 6 in

the 8-week model) (Figure 6). Histology was used to assess autophagy and cardiac fibrosis

parameters, as described in detail in the Materials and Methods section. Echocardiography

was performed at 4 or 8 weeks to evaluate functional parameters such as ejection fraction

(EF%), fractional shortening (FS%), heart rate (HR), and left ventricular posterior wall

thickness diameter (LVPWd). All rats with MI were examined by an echocardiographist to

confirm the results.

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Results

Fibroblast phenotype and inhibition of autophagy in the post-MI heart

We utilized Western blotting to determine changes in protein levels of the autophagy

markers LC-3I and LC-3II, with β-tubulin as a control for total protein loading. At 4 weeks

post-MI, although we observed a trend towards increased accumulation of the autophagy

marker LC-3I in untreated (no drugs; ND) viable and scar regions of the heart, the differences

were not significant (Figure 18A). No notable changes in LC-3II accumulation were observed

in untreated hearts post-MI. However, in CQ-treated hearts, there was a modest (but non-

significant) trend towards increased LC-3I in the viable region and a significant (P ≤ 0.05)

increase in LC-3I in the scar region, compared to sham-operated CQ-treated controls (Figure

18A). We observed a trend towards an increase in LC-3II in viable and scar regions of post-

MI CQ-treated hearts, though this change was not significant (Figure 18A). When comparing

ND and CQ-treated groups, we observed that CQ treatment caused increased accumulation of

LC-3I in post-MI viable and scar regions. In summary these results show increased

autophagy in the infarct scar vs control. CQ treatment in scar region led to significant LC-3II

accumulation which is associated with inhibition of autophagy.

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A

113

B

C

B

C

114

Figure 18: LC-3 lipidation as a marker of autophagy in 4 week untreated and

chloroquine-treated post-MI rat hearts (control, myocardium remote to the scar and the

infarct scar). Panel A. Western blot for autophagy markers LC-3I and LC-3II in untreated

(no drugs; ND) and chloroquine (CQ)-treated hearts. Levels of LC-3I trended toward increase

in the post-MI scar region of untreated (no drug; ND) hearts. No change in LC-3II levels was

observed between untreated sham-operated and MI hearts. CQ treatment (40mg/kg-1

/day for 2

weeks beginning at 1 week post-MI) induced significantly higher levels of LC-3I in post-MI

scar regions, but not viable regions. CQ treatment produced a trend towards increased levels

of LC-3II in post-MI viable and scar regions, though levels did not differ significantly. ND:

no drugs, #compared to sham-operated, P < 0.05.Significance was determined using one-way

ANOVA followed by a Student-Newman-Keuls post-hoc test.

D

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Panel B. Western blot for LC-3I and LC-3II in sham-operated untreated vs. CQ-treated

hearts. There was no significant difference in autophagy marker levels in sham-operated

hearts in untreated vs. CQ-treated groups. Panel C. Western blot for LC-3I and LC-3II in the

viable region of post-MI hearts, comparing untreated vs. CQ-treated groups. There was a

significant increase in LC-3I expression in the viable region of post-MI hearts treated with

CQ, compared to untreated hearts(*P<0.05 compared to no drugs; ND).Significance was

determined using student-t test. No changes were observed in LC-3II levels with CQ

treatment. Panel D. Western blot for LC-3I and LC-3II in the scar region of post-MI hearts,

comparing untreated vs. CQ-treated groups. No changes were observed in LC-3I levels for

untreated and CQ-treated hearts. There was a significant increase in LC-3II expression in the

scar region of post-MI hearts treated with CQ, compared to untreated scar controls. (#P<

0.05compared to no drugs, data is mean ± SEM, n=3-6 distinct cardiac samples per

group).Significance was determined using student-t test.

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To determine the effect of MI and CQ treatment on myofibroblast presence in rat

hearts at 4 weeks, we performed Western blotting analysis for the key myofibroblast marker

α-SMA. In untreated hearts, MI was associated with increased α-SMA protein levels in viable

tissue, though this difference was not statistically significant. However, MI is linked to a

marked and significant (P< 0.05) increase in α-SMA levels in scar tissue vs controls (Figure

19). These results indicate an increase in cardiac fibroblast activation at 4 weeks post-MI in

the infarct scar of these rats. CQ treatment was effective in significantly (P< 0.05) reducing

α-SMA expression in both the viable and scar regions of post-MI hearts, compared to

untreated controls for these regions (Figure 19).

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Figure 19: -SMA marker levels in 4 week untreated and chloroquine-treated post-MI

rat hearts.Western blot analysis of α-SMA (marker of activated fibroblasts)from left

ventricular tissue lysates of sham-operated and MI rat hearts, either untreated or treated with

chloroquine (CQ; 40mg/kg-1

/day for 2 weeks beginning at 1 week post-MI). α-SMA

expression in untreated and CQ-treated sham hearts was low. In scar regions of post-MI

hearts, α-SMA was significantly increased, compared to viable post-MI and sham-operated

hearts. Treatment with CQ significantly reduced α-SMA levels in these regions, as compared

to the same regions with no drug (ND). (*compared to sham ND; #compared to MI viable

ND; фcompared to MI viable ND;

ccompared to MI scar ND; P< 0.05 for all comparisons).

118

Data is mean ± SEM (n=3-6 distinct cardiac samples per group). ND: No drugs. Significance

was determined using one-way ANOVA followed by a Student-Newman-Keuls post-hoc test.

119

To complement these findings in left ventricular lysates, we used immunostaining to

examine autophagy (via LC-3β accumulation) and fibroblast activation (α-SMA staining) in

cardiac tissues at 4 weeks in sham-operated and MI groups with and without CQ treatment.

Immunofluorescence staining for LC-3β staining was increased in post-MI hearts with and

without CQ treatment (Figure 20). We also observed an increase in α-SMA staining in post-

MI hearts, compared to sham-operated controls, which was apparently diminished with CQ

treatment post-MI. These results confirm that elevated autophagy accompanies accumulation

of α-SMA untreated infarct scar. CQ treatment was associated with reduced expression of α-

SMA in the left ventricle at 4 weeks post-MI. Thus treatment with CQ in 4 week post-MI rat

hearts is associated with reduction of fibroblast activation when compared to the untreated

MI rat hearts.

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Figure 20: Immunofluorescence staining of post-MI and sham-operated hearts at 4

weeks, with and without chloroquine treatment. Basal levels of LC-3β (autophagy

marker) and α-SMA (myofibroblast marker) were observed in sham-operated controls, with

and without CQ, are increased in post-MI ventricles. (CQ; 40mg/kg-1

/day for 3 weeks

beginning at 1 week post-MI). Magnification 10X.

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As the reduction ofα-SMA expression in CQ treated 4 week-post MI rats was

significantly reduced, it is reasonable to explore the concomitant effect of CQ treatment on

collagen accumulation in infarcted myocardium using Masson’s trichrome imaging and

hydroxyproline assay. While increased collagen content was present in the 4 week post MI

rat heart sections vs. controls, we found that CQ treatment of these animals did not affect

collagen levels post-MI

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Figure 21: Masson’s trichrome staining for collagen content in 4 week post-infarct left

ventricles, with and without chloroquine treatment. Blue staining in control and

experimental sections above reveals collagen deposition. At 4 weeks after infarction, collagen

staining is increased in intensity and density of staining, and is unchanged with CQ treatment

vs. untreated controls. Magnification 4X.

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To quantify and validate our Masson’s trichorme stain experiements, we assayed

hydroxyproline content across various groups. The helical structure of collagen is due to the

post-translational hydroxylation of the prolines present in the collagen. Thus presence of

hydroxyproline is restricted to fibrillar collagens and elastin. Due to the low content of elastin

proteins, the precedent in the literature is to use hydroxyproline content to infer myocardial

fibrillar collagen content [294]. Our results with this second assay paralleled those noted

inMasson’s staining experiments. That is, we found increased collagen accumulation found in

4 week post-MI untreated animal vs control. There was no significant reduction found in our

CQ treated MI groups vs untreated MI groups. This set of data revealed that the CQ treatment

is linked to deactivation of fibroblasts, but not with any change in collagen accumulation.

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Figure 22: Hydroxyproline collagen quantification in 4 week post MI cardiac tissue:

Histographical representation of the hydroxyproline content in 4 week post MI rat hearts

indicate increased collagen content in the MI untreated groups when compared to untreated

sham. We observed no significant difference in myocardial hydroxyproline content of the

MI+ CQ treated samples vs. control. Values are expressed as mean of 4 -6 per group. (*

P<0.05 experimental vs. controls). Significance was determined using one-way ANOVA

followed by a Student-Newman-Keuls post-hoc test.

*

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Similar to in the 4-week study group, we assessed expression for autophagy and

myofibroblast markers in 8-week study group hearts post-MI for untreated and CQ-treated

animal tissue lysates. Western blot analysis for LC-3I and LC-3II show a trend (but not

significant) toward increased expression of LC-3II in MI scar tissue, compared to MI viable

and sham-operated, in untreated hearts (Figure 23). CQ-treated groups show a similar trend

(non-significant) to increased expression of both autophagy markers (Figure 23). Our analysis

of α-SMA expression showed similar results. In untreated hearts, α-SMA levels were

significantly increased in the MI scar, but not in the viable region, compared to sham-

operated controls (Figure 24). Treatment with CQ appeared to show a trend (non significant)

towards decreased α-SMA levels in the MI scar region (Figure 24). Immunofluorescence

staining for 8-week study group hearts post-MI with and without CQ also support the

Western blot data, showing some degree of decrease in α-SMA staining with CQ treatment

post-MI (Figure 25).

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Figure 23: LC-3β lipidation as a marker of autophagy autophagy in 8-week untreated

and chloroquine-treated rat hearts post-MI. Expression of autophagy markers LC-3I and

LC-3II in untreated (no drugs; ND) and chloroquine (CQ) treated hearts show trending

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increase in LC-3II levels in the post-MI scar region of untreated (no drug; ND) heart and CQ

treated heats when compared to sham-operated treated and untreated groups respectively.

However this did not reach significance. Data are expressed as mean ± SEM (n=3 - 4). ND:

No drugs.

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Figure 24: α-SMA expression in 8-week post-MI hearts treated with chloroquine. α-

SMA expression was assessed in CQ treated and untreated post-MI hearts after at end of 8

week post-MI. We saw a significant increase in α-SMA expression in the infarct scaring the

non-treated groups when compared to sham-operated ND group (*compared to sham ND). In

contrast, no significant similar increase ofα-SMA expression was observed in the MI+ Scar

CQ treated vs. the untreated group of same region. We interpret this finding to show that

autophagy is suppressed in the CQ treated infarct scar samples. Data is the mean ± SEM of

n=3-4 hearts). ND: No drugs. Significance was determined using one-way ANOVA followed

by a Student-Newman-Keuls post-hoc test.

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Figure 25: Immunofluorescence staining of 8 week post-MI hearts with and without

chloroquine. LC-3β (autophagy marker) and α-SMA (myofibroblast marker) were observed

in sham-operated controls, with and without CQ, with increase in post-MI ventricles. (CQ;

40mg/kg/day for 3 weeks beginning at 5 week post-MI). Mgnification 10X.

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Assessment of Biochemical Parameters for Cardiac, Liver and Kidney Damage

The enzymes routinely measured clinically to diagnose and monitor myocardial

infarction include lactate dehydrogenase (LDH) in the early stages following MI. Increased

LDH long-term may indicate damage to various tissues, including the heart, liver, kidney,

muscle, lungs, and erythrocytes [333, 334]. To diagnose organ damage, biochemical assays

are used to measure serum levels of creatinine for kidney damage [335], and aspartate amino

transferase (AST)and aspartate transaminase (ALT) for liver damage [336]. There was no

difference in the LDH or creatinine levels in any of the groups at 4 and 8 week post-MI when

compared to sham-operated controls, suggesting that the drug has no effects of widespread

tissue or kidney-specific damage. We observed a trend towards elevation in AST and ALT

levels at 4 and 8 weeks in the CQ treated MI groups (Figure 26), indicating a possibility of

low-level sustained damage to the liver following CQ treatment post-MI.

132

A

* *

133

B

134

Figure 26: Biochemical parameters for cardiac, kidney and liver damage in 4 and 8

week post-MI rat hearts with and without chloroquine treatment. Panel A. At 4 weeks,

no significant differences were observed in serum markers for wide spread tissue damage

(lactate dehydrogenase; LDH) nor damage to the kidney (creatinine). There was a significant

increase in serum marker of liver (aspartate amino transferase (AST) and aspartate

transaminase (ALT).(* P< 0.05 MI vs. MI + CQ). Significance was determined using one-

way ANOVA followed by a Student-Newman-Keuls post-hoc test. Panel B. At 8 weeks,

there are still no significant increases in these damage markers, though again there is a trend

towards increased AST and ALT in the CQ-treated post-MI group. (CQ; 40 mg/kg-

1/day).Data expressed as mean ± SEM (n=4-6).

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Functional Assessment Following Myocardial Infarction and Chloroquine Treatment

To determine whether CQ treatment of post-MI animals translated to improvements in

cardiac functional changes, we assessed cardiac functional parameters using

echocardiography. We confirmed via biochemical assay that our 40mg/kg-1

/day CQ drug

regime did not invoke tissue damage (Figure 26).Using non-invasive transthoracic

echocardiography (TTE), we obtained the M-mode echocardiography images and evaluated

the standard functional parameters of cardiac function including injection fraction (EF),

fractional shortening (FS), left ventricular posterior wall diameter (LVWPd) and heart rate

(HR). All the parameters were monitored in the second, third and fourth week in the 4-week

study group, and during the fifth, sixth and eighth week in the 8-week study group. In the 8-

week study group, we observed significant reductions in ejection fraction to approximately

60% in post-MI animals, compared to sham-operated controls, remaining consistent

throughout the 2-4 week period. Treatment with CQ was not associated with any change in

the ejection fraction in infarcted animals, which remained significantly lower than in sham-

operated + CQ animals and at similar levels as untreated post-MI groups, around the 60%

mark (Figure 27A). We also observed significantly decreased fractional shortening,

indicating reduced contractility, from~50% in sham-operated-operated controls (with and

without CQ) to less than 30% in post-MI animals, either with or without CQ treatment

(Figure 27A). In the 8-week study group, we observed similar results, with ejection fraction

being reduced by about 30% in post-MI animals with or without CQ. Similarly, fractional

shortening was reduced to less than 30% in all post-MI animals, again regardless of CQ

treatment (Figure 28B). Heart rate and LVPWd were highly variable in the 4-week treatment

group, but there were no significant differences between any of the treatment or control

groups (Figure 28A). Similarly, we found no difference in these parameters in the 8-week

study group (Figure 28B).

136

A

137

B

138

Figure 27: Ejection Fraction and Fractional Shortening in Post-MI Rats. M-Mode

transthoracic echocardiography (TTE) was performed at 2, 3 and 4 weeks (for 4-week study

groups) or 5, 6, and 8 weeks (for 8-week study groups) after the LAD- ligation surgery to

assess ejection fraction (EF) and fractional shortening (FS). Panel A. In the 4 week study

group, EF and FS were both significantly reduced following MI, with or without CQ

treatment. Panel B. In the 8 week study group, EF and FS were significantly reduced post-MI

regardless of CQ treatment. No progressive decrease in function was observed over the

duration of TTE examinations. (*compared to sham-operated; #compared to sham-operated +

CQ; P < 0.05 for all comparisons). Data is mean ± SEM (n=4-6). Significance was

determined using one-way ANOVA followed by a Student-Newman-Keuls post-hoc test.

139

A

140

B

141

Figure 28: Heart Rate and Left Ventricular Posterior Wall Diameter in Post-MI Rats.

M-Mode transthoracic echocardiography (TTE) was performed at 2, 3 and 4 weeks (for 4

week study groups) or 5, 6, and 8 weeks (for 8 week study groups) after the LAD- ligation

surgery to assess heart rate (HR) and left ventricular posterior wall diameter (LVPWd). In

both 4 week (Panel A) and 8week (Panel B) study groups, HR and LVPWd did not change

following MI, with or without chloroquine treatment (40 mg/kg-1

/day). Data is mean ± SEM

(n=4-6).

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Discussion

We have investigated the role of autophagy in cardiac fibrosis and its impact on heart

function using an in vivo experimental model of MI in combination with CQ treatment. We

utilized Western blotting and immunofluorescence to assess changes in protein levels for

autophagy and myofibroblast markers in the 4-week post-MI model. We observed significant

increases in LC-3I (in the viable region) and LC-3II (in the scar region) in post-MI animals

treated with CQ (Figure 18, Panels C and D) vs. untreated post-MI hearts, as well as

increased LC-3I in the scar region post-MI compared to sham-operated controls, both of

which were treated with CQ (Figure 18A). Controversy regarding interpretation of LC-3 data

from Western blots exists in the literature [337], and it is suggested that this lies within the

approach used to calculate lipidation ratios [338]. In addition, LC-3II may be degraded by the

autophagolysosome, and needs to be measured against an internal control, such as beta-

tubulin. Increased LC-3II levels are associated with increased autophagosome formation (or

decreased turnover, if trafficking to lysosomes is delayed or there is reduced fusion or

impaired proteolytic activity of lysosomes). However, in the presence of inhibitors such as

Baf-A1 and CQ, increased LC-3II levels are observed, due to inhibition of lysosomal

proteases and impaired degradation of lysosomal contents [155]. Failure of these inhibitors to

enhance LC-3II levels would indicate defects earlier within autophagy, prior to the fusion

events wherein these inhibitors have their effects [338]. In 4 week post-MI experimental

groups, autophagy was significantly elevated in the infarct scar, and was inhibited with CQ

treatment(Figure 18). However this result was not associated with any change in collagen

staining of post-MI hearts, nor in their hydroxyproline content with CQ treatment. Taken

together, these results indicate that CQ treatment diminished α-SMA and inhibited

autophagic flux in treated hearts following MI at 4 weeks, but that these changes were not

linked to any effect on collagen deposition. We suggest that this scenario exists because of

143

the timing of the experiment: that is, with initiation of CQ treatment, fibrosis and collagen

deposition are already well underway, along with reduced EF in experimental animals. Also,

it is known that the biological half-life of collagen is 80-120 days [117], which means that

lag-time of any anti-fibrotic treatment is significant. While CQ may restrict activation of

fibroblasts, the window for the duration of treatment is too narrow to be asscociated with

reduction of collagens. We suggest that an earlier administration of CQ post-MI could have

restricted the formation of collagen and inhibited fibrosis.

The role of autophagy in cardiac pathology remains controversial, as some reports

emphasize a protective role for autophagy in the heart, whereas other reports emphasize its

negative effects on cardiac function. Liu et al. performed in vitro and in vivo autophagy

experiments in cardiac fibroblasts and C5BL/6 mice, respectively, which demonstrated a

protective role for autophagy against lead toxicity [104]. When 3MA was used to inhibit

autophagy, this group observed enhanced lead toxicity in cardiac fibroblasts, resulting in cell

death [339]. Another study by Liu et al. in 2016 used ANGII treatment both in vitro and in

vivo to induce cardiac fibrosis. In this study, the researchers found that enhancing autophagy

(via rapamycin) decreased upregulation of collagen type I and fibronectin expression in rat

cardiac fibroblasts, and inhibition with CQ attenuated these effects. In vivo, rapamycin in

combination with angiotensin II infusion ameliorated both cardiac fibrosis and dysfunction,

while inhibition with CQ worsened these ANGII-mediated effects [340]. Xie et al. found that

chronic metformin treatment of type I diabetic mice enhanced autophagy and improved

cardiac function, indicating a protective role for autophagy in this model [271].

Other reports have provided evidence to underscore autophagy as a maladaptive

response and have demonstrated therapeutic potential for autophagy inhibitors. Xu et al.

found that in a diabetic mouse model, wild type diabetic hearts had increased levels of

oxidative stress, interstitial fibrosis, myocyte apoptosis, and reduced cardiac function

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compared to heterozygous Beclin-/- and Atg16-/- animals [341]. In that study, Beclin-1

rescue via over-expression restored autophagy and worsened cardiac function, demonstrating

an adverse role for autophagy in the hearts of diabetic mice. Similarly, a report of transverse

aortic constriction (TAC)-induced model of pressure overload, TAC increased cardiac

autophagy to elevated levels for durations reaching 3 weeks. Further enhancement of

autophagy by Beclin-1 overexpression further heightened autophagic activity and increased

adverse remodeling in pressure-overloaded hearts [245]. The variable effects of autophagy in

different models of cardiac pathology indicate a complex role for this process in the heart. It

is also pointed out that multiple cell types exist in the heart and that autophagy may occur in

myocytes, endothelial cells, smooth muscle cells, and fibroblasts, and that some of the

aforementioned studies did not distinguish among them. In our current in vivo work, the

distinction between these cell types was also difficult to resolve. Other limitations of our in

vivo studies may be present in the time of treatment of the post-MI animals with CQ, as the

cardiac collagen levels (4-OH levels) in treated vs. untreated samples was similar between

groups. As mentioned above, we speculate that earlier CQ administration post-MI in our

experimental design may have effected collagen accumulation. Finally we focused the current

study on detection of systolic dysfunction in treated and untreated damaged hearts. This point

could be argued to be a design omission within the experimental design as diastolic left

ventricular (LV) dysfunction is known to be primarily associated with heart failure with

preserved ejection fraction (HFpEF) [342], and while we recognize that this is important

feature of fibrosis, we were not able to fully assess putative subtle changes in cardiac

diastolic function. This issue may have limited our ability to link cardiac filling dysfunction

between treated and untreated groups in damaged hearts, even in the presence of unaltered

global cardiac collagen levels among groups.

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Other studies using animal models of MI have highlighted the role of autophagy in

post-MI remodeling and cardiac fibrosis, and points to the involvement of temporal changes

in autophagic flux in the injured heart. In 2014,a study by Wu et al. revealed that in mouse

subjected to LAD-induced MI, increased autophagic induction occurred in the acute phase of

MI (0.5-3 days) and was subsequently reduced in the latter phase of MI (5-21 days). One day

post-MI, treatment with rapamycin reduced post-MI remodeling and dysfunction, whereas

inhibition with 3MA worsened these outcomes [290]. Zhang et al. found that berberine-

induced increases in LC3-βII and beclin-1 expression were cardioprotective at 4 weeks post-

MI, reducing interstitial fibrosis and adverse remodeling and improving ejection fraction and

fractional shortening [343]. These studies underscore the importance of autophagy in the

pathogenesis of heart failure as it relates to cardiac fibrosis.

Although the link between autophagy and cardiac fibrosis is still relatively

unexplored, reports in lung, liver and kidney reveal a positive correlation between autophagy

and fibrosis [310, 344] and advocate for anti-autophagic molecules as novel therapeutics.

This is also supported by our earlier work in human atrial fibroblasts demonstrating that

TGF-β-mediated elevation of profibrotic markers collagen type I and fibronectin was

associated with increased autophagic flux [256]. In this study, we also demonstrated

concomitant increases in levels of protein markers of fibrosis, autophagy, and Smad2

phosphorylation (indicating increased TGF-β signaling) in whole scar tissue lysates from the

hearts of post-MI rats at 2 and 4 weeks. Atg5-knockout mouse embryonic fibroblasts also

showed decreased fibronectin synthesis [256]. While our in vitro (Chapter 1) results are

supportive of the involvement of autophagy inhibition in attenuating fibroblast mediated

profibrotic events, our in vivo data (Chapter 2) do not support the argument for the

development of therapeutic agents that influence autophagy in cardiac fibroblasts. We

suggest that addressing the autophagy/cardiac fibrosis hypothesis remains comprehensive in

146

the context of influencing cardiac fibrosis in vivo, but with a refined study using an early time

point for CQ administration or, alternatively, other novel autophagy inhibitory agents with

greater specificity.

Morphological and structural changes are not clinically significant without

corresponding significant changes in cardiac function that leads to improved patient

outcomes. We performed echocardiography in all animal groups, a clinically relevant tool to

assess cardiac function. The procedure is especially useful due to its non-invasiveness and

reflects ventricular dysfunction and hemodynamic alterations resulting from cardiac

pathologies [132]. As expected, MI resulted in decreased functional changes as early as 2

weeks post-MI compared to sham-operated controls, namely ejection fraction and fractional

shortening (Figure 27), which are indicators of pumping efficiency and contractility,

respectively [130, 345]. Notably, these parameters were not altered with CQ treatment of

damaged hearts in either the 4- or 8-week study groups. Similarly, heart rate of post-MI rats

did not change significantly with CQ treatment over the time durations examined in this study

(Figure 28). In the 4week study group, we saw no change in left ventricular posterior wall

diameter (LVPWd), though there was a modest trend towards decreased LVPWd over time

(eg, 5 to 8 weeks) with CQ treatment (Figure 28).The lack of functional changes and lack of

any change in collagen content in treated post-MI hearts vs. controls may be due to

insufficient dosage. Our study used 40 mg/kg/day, but other in vivo studies used from 100 to

500 mg/kg/day[346]. We chose to treat animals with a lower dose with respect to the

possibility of generation of side-effects. In support of this presumption, that serum markers of

tissue damage were not significantly increased with CQ treatment in the current study (Figure

26).Physiologically relevant dosages for CQ will likely differ between species, considering

differences in metabolism and heart size, among a host of other factors affecting the response

to CQ and potential side effects.

147

In light of the above observations, our study does support the central hypothesis of

attenuating the activation of fibroblast, however it does not support that low-dose CQ

treatment in a rat model of MI could reverse or attenuate adverse cardiac remodeling. This

notwithstanding, our in vitro work provides supportive evidence for future research to

address autophagy inhibition in treatment of cardiac fibrosis in vivo.

148

General Discussion

Interest in cardiac fibrosis as a primary cause amongst the pathogenesis of heart

failure is burgeoning., following MI [11]. Cardiac fibrosis is characterized by the chronic

accumulation of collagen via the persistence of activated myofibroblasts in the infarcted and

distal regions of the heart, long after wound healing associated with scar formation is

complete [347-349]. In recent years, autophagy has been suggested to regulate cardiac

remodeling, and dysregulated levels of autophagy are implicated in various cardiac

pathologies including pressure overload-induced hypertrophy and fibrosis [258, 340, 343].

Gao et al. reported elevated autophagy with in a diabetes-induced cardiac fibrosis rat model

[350]. A role for autophagy in fibrosis of other organs, including the lungs and kidney has

already elucidated [167, 351-353]. However, the relationship of cardiac fibroblast activation

and autophagy in the context of myocardial infarction and heart failure is still unexplored. In

the current study, we have focused on understanding the relationship between autophagy and

fibroblast activation using pharmacological inhibition of autophagy in unpassaged (P0)

primary cardiac fibroblasts (in vitro) and in an in vivo experimental model of MI 4 and 8

weeks post-MI.

In our in vitro study, when unpassaged primary cardiac fibroblasts were exposed to

mechanical stress, we found enhanced activation of fibroblasts with concomitant conversion

to myofibroblasts. This was demonstrated by increased α-SMA and ED-A FN protein levels

as assessed by Western blotting and immunofluorescence (Figures 7 and 11, respectively).

Concomitant to this, we found elevated expression of LC-3βII, which is an important

molecular marker of autophagy (Figure 8). Several reports are available where autophagy

inhibitors have been used to attenuate fibrosis, including well-known inhibitors 3MA and CQ

[343, 354], though one report indicated worsening of ANGII-induced fibrosis following CQ

treatment [340], and there are no studies examining the effects of Baf-A1 on cardiac fibrosis.

149

In our experiments, these inhibitors successfully inhibited autophagy in the in vitro model,

confirmed by both Western blotting and TEM (Figures 8 and 9). These inhibitors, in a dose-

and time-dependent manner, caused reduced activation of P0 rat cardiac fibroblasts to the

myofibroblast phenotype, indicated by significant reductions in α-SMA and ED-A FN

proteins levels compared to controls (Figure 10). Immunofluorescence staining of unpassaged

cardiac fibroblasts treated with Baf-A1, 3MA, or CQ showed concomitant decreases in LC-

3II staining and α-SMA stress fiber formation (Figure 11). Functional myofibroblast

functions, including motility and contractility, were also reduced with autophagy inhibition

(Figures 12 and 13). These lines of evidence provide strong support for the role of autophagy

in the fibroblast activation and the efficacy of autophagy inhibitors in preventing the

myofibroblast phenotype.

Among the three inhibitory drugs, CQ appeared to be the most effective inhibitor of

fibroblast activation, thus we utilized CQ to further understand the mechanism underlying the

relationship between autophagy inhibition and reduced activation of cardiac fibroblasts to

myofibroblasts. The MAPK signaling pathway has been previously implicated in autophagy

in the contexts of cardiac hypertrophy and fibrosis [287]. Our preliminary study on MAPK

signaling proteins such as p38 and ERK showed significant changes with p38 MAPK

phosphorylation, however we did not see any change with ERK (p44/42) with time. Thus,

we monitored p38 MAPK activation by examining the ratio of phosphorylated p38 MAPK

protein to total p38 MAPK protein (via Western blotting) in the presence and absence of

autophagy inhibitors. In the absence of autophagy inhibitors, we found a time-dependent

decrease in p38 MAPK activation with plating on a stiff substrate (Figure 14). This result is

as expected, as decreased p38 MAPK signaling has shown to be associated with the

conversion of fibroblasts to myofibroblasts [101]. The finding that treatment with CQ

increased p38 MAPK activation agrees with our findings that autophagy inhibition prevents

150

fibroblast activation (Figure 14). Co-inhibition of MAPK signaling and autophagy did not

demonstrate any cooperative effects, though this strategy remained effective in attenuating

activation of P0 cardiac fibroblasts (Figure 15). Nonetheless, these findings demonstrate a

putative link between MAPK signaling and autophagy inhibition.

After successfully establishing a link between autophagy and cardiac fibroblast

activation, as well as attenuation of these events with CQ treatment of primary fibroblasts, we

tested the efficacy of CQ in an in vivo experimental model of MI. We confirmed an increase

in myocardial collagen deposition (Figure 21 and 22) as well as α-SMA expression (Figure

19) in untreated experimental animals, indicating increased myofibroblast presence,

concomitant with increased autophagic flux (Figure 18) at both 4 post-MI. These results

agree with previous findings linking increased autophagy and the development of cardiac

fibrosis and subsequent heart failure [255, 290, 355]. In 4 week groups treated with low-dose

CQ for 3 weeks prior to tissue harvesting, α-SMA expression and was attenuated with

inhibition of autophagy in agreement with our findings in vitro. However, when we

compared collagen content and carried out functional assessment of heart was performed by

echocardiography in MI and CQ + MI groups we observed no effect with low-dose CQ

treatment of experimental hearts vs. non-treated controls. Indeed, ejection fraction, heart rate,

and fractional shortening were compromised in MI groups (compared to sham-operated

controls) and were unaltered with CQ treatment (Figure 27 and 28). However, serum markers

for liver, kidney, and cardiac damage were significantly reduced in the CQ + MI group

(compared to MI group), though not to the same level as the control group (Figure 26).As

changes in these markers precede functional alterations in the heart, the discrepancy between

serum marker levels and echocardiography results may be due to the timing of low-dose CQ

treatment. Thus, further optimization of the study timeline, such as commencing CQ

151

treatment earlier, or extending treatment to 8 weeks, may prove fruitful in observing similar

effects between marker levels and functional assessment.

152

Study Conclusions

Overall, the findings of this study lead us to three major conclusions, which are:

1. Plating of primary, unpassaged P0 cardiac fibroblasts on stiff substrates induces the

activation of fibroblasts, and in parallel their conversion to the myofibroblast phenotype,

as indicated by elevated expression of myofibroblast markers.

2. Activation of cardiac fibroblasts is associated with decreased p38 MAPK activation and

increased autophagic flux in myofibroblasts, indicated by the markers LC-3B and p62, as

well as autophagosome formation (assessed via TEM), both in vitro and in vivo.

3. Inhibition of autophagy prevents activation of fibroblasts in vitro and ameliorates -SMA

elevation post-MI in vivo. However the application of relatively low-dose CQ to post-MI

animals in vivo was not associated with any change in systolic cardiac function nor in

cardiac hydroxyproline (collagen) content. Future experiments with different antifibrotic

agents and/or timing of CQ treatment to earlier points after MI are likely warranted.

Thus, the in vitro segment (Chapter 1) of our study provides supportive evidence for

further examination of autophagy inhibition as a therapeutic strategy for attenuating

activation of cardiac fibroblasts and ensuing profibrotic events mediated by cardiac

myofibroblasts. The results of our in vivo study does restrict the phenotypic shift in the 4

week model, but not support the use of relatively low-dose CQ as an inhibitor of post-MI

cardiac fibrosis. Limitations in our in vivo experimental design may have impacted on the de

facto efficacy of our anti-autophagic treatment.

153

Future Directions

The in vitro findings above provide support for the induction of autophagy in driving the

activation of cardiac fibroblasts. However, questions still remain regarding the efficacy of in

vivo autophagy inhibition in preventing or reversing cardiac fibrosis and the specific cellular

mechanisms involved.

1. Pan-specific or inducible genetic knock-out mouse models to exploit the autophagic

pathway is a possible avenue to be explored, in combination with MI induction or

TAC-induced pressure overload. Specifically the knock-out of key autophagic genes

such as ATG7 might be used to assess the specificity of autophagy suppression (both

in vitro and in vivo). This approach could be applied to bypass the usual issues

associated with drug delivery and efficacy, particularly in lieu of the lack of a highly

effective small-molecule inhibitor of autophagy.

2. Additionally, the role of autophagy in cardiac fibrosis can be further understood by

targeting other important signaling pathways such as mTOR and PI3K. To elucidate

the role of mTOR and PI3K pathways, one could use mTOR specific and mTOR and

PI3K dual specificity inhibitors, either alone or in combination [356].

3. Fibroblasts are mechanically responsive cells. In the context of our work, there may

be relevant mechanical signaling pathways involved in triggering autophagy and

activation of fibroblasts. The use of various mechanical stimuli such as culture of

fibroblasts subjected to repeated and controlled mechanical stretch, and co-plating

with other cell types like cardiomyocytes and/or on various substrates would allow

cell-cell and cell-matrix interactions, which could then be studied in terms of

autophagy and fibroblast activation.

4. The question of the precise mechanism(s) of autophagy in fibroblast activation is far

from clear. We speculate that autophagy affects MAPK (p38) signaling, but we know

154

that other molecules may influence autophagy and also have anti-fibrotic properties.

Our lab has recently shown that Ski effectively inhibits autophagy in cardiac

myofibroblasts [357], it will be important to understand which, if any, autophagy-

related genes are affected. To this end, microarray data in myofibroblasts treated with

autophagy inhibitors may provide a starting point to determine the specific genes

involved in the autophagy-fibroblast activation relationship.

5. The effects of enhanced autophagy using rapamycin with such models can also be

studied to examine the integrity of the relationship between autophagy and cardiac

fibrosis. Although other studies have examined cardiomyocyte autophagy in cardiac

fibrosis, it will be important to understand the effects of this agent in myofibroblasts

and to assess for effects on profibrotic events.

6. Altered timing of administration of inhibitors of fibroblast autophagy: If the lack of

efficacy of CQ – mediated autophagic inhibition in vivo are due to alterations in the

activation of fibroblasts, a time course study to address earlier phases (eg, earlier than

4 weeks of post-MI wound healing) will need to be considered. For example,

administering CQ within 2-5 days post-MI could not only limit the activation of

fibroblast but it might also restrict the collagen production and fibrosis. This will also

help to better understand the initial phases of post-MI wound healing and scar

formation in the context of autophagy activation. In addition to a comprehensive time

course study, the use of different doses of CQ will also allow us to determine a dose-

response. In addition to CQ, the use of newer, more potent autophagy inhibitors such

as Lys01 and Lys05 may provide a clearer understanding of the potential for these

drugs as anti-fibrotics in real-world applications. If efficacy can be demonstrated with

optimized timing and drug usage/dosage in experimental MI, this will help to

determine the putative effectiveness in other models of MI and pressure overload. It

155

will also be interesting to examine the effects of autophagy inhibition in fibrosis of

other tissues and organs, particularly those where fibroblast activation is well-studied,

such as dermal models. Ultimately, the use of statistical modelling and regression

analysis in a review of peer-reviewed publications on the subject of autophagy

inhibition in cardiac remodeling and fibrosis in animal models would provide proof-of

concept evidence to bring these methods to the clinical realm.

7. A perfenidone study: Perfenidone is a late generation antifibrotic drug used to treat

idiopathic pulmonary fibrosis (IPF). This drug has been shown to have anti-fibrotic,

anti-inflammatory and antioxidant properties. In vitro studies have shown that it limits

TGF-β stimulated collagen production by downregulating theCol1 mRNA and

fibroblast proliferation. It also reduces production of pro-inflammatory cytokines and

free radicals[358].It would be interesting to determine whether combination therapy

using CQ as autophagy inhibitor and perfenidone as antifibrotic drug could have

beneficial effects on cardiac fibrosis. Our study CQ shows that CQ is an efficient

blocker of myofibroblast phenotype. A combination both drugs could be a new area of

research in treatment of cardiac fibrosis.

156

Limitations of this Study

This study is focused on putative changes to the autophagic state in heart after MI in rat heart

model of post-MI remodeling. The purpose is to definitively study the role of autophagy in

the activation of cardiac fibroblasts in vivo and in the second phase of the study, of cardiac

fibrosis after the induction of MI. While diastolic dysfunction and cardiac hypertrophy are

important features of heart failure, per se, in post-MI cardiovascular disease, our study does

not address the occurrence of heart failure directly. Rather, we have focused on fibroblast

activation and fibrosis and for this purpose we evaluated the protein markers for phenotype

and collagen deposition in our post-MI rat model. Cardiac performance was analyzed using

M-mode echocardiography (rather than by direct intubation of the left ventricle with a force

transducing catheter), which provides data giving information on systolic dysfunction. Thus,

the limitations of our in vivo study are that we did not monitor parameters of diastolic

dysfunction and heart failure such as E/A ratio (ratio of peak velocity flow in early diastole

(the E wave) to peak velocity flow in late diastole caused by atrial contraction the A wave),

LVEDP, and especially liver, abdominal and lung ascites (wet/dry ratio). In retrospect, our

study would have been more comprehensive had we provided data on cardiac hypertrophy by

calculating heart weight/tibia length ratio.

157

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