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Host protein manipulation as a mechanism in viral cardiomyopathy Wong, Tse Yuan 2012

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HOST PROTEIN MANIPULATION AS A MECHANISM IN VIRAL CARDIOMYOPATHY  by  Tse Yuan Wong  B.Sc., Simon Fraser University, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  The Faculty of Graduate Studies  (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)  December 2012  © Tse Yuan Wong, 2012  Abstract Viral myocarditis, the inflammation of myocardium initiated by viral infection, is an important cause of mortality in neonates and children. In addition, it is a precursor to dilated cardiomyopathy (DCM). To date, no effective therapy is available for viral myocarditis/DCM. Coxsackievirus B3 (CVB3) is an important human pathogen of viral myocarditis. Extensive research efforts on CVB3 have broadened our understanding of the virus-host protein interactions. However, the pathogenesis of coxsackievirusinduced myocarditis is not fully understood. The objective of this dissertation is to explore the role of host protein manipulation in coxsackieviral replication and pathogenicity. My hypotheses are that (1) coxsackievirus hijacks host’s cellular autophagy mechanism to facilitate its own replication; and (2) the serum response factor (SRF) is cleaved by viral protease 2A during coxsackievirus infection and contributes to impaired myocardial function and progression to DCM. For project 1, I demonstrated that CVB3 manipulates the host autophagy pathway to supplement viral replication. Autophagy is an evolutionary conserved homeostatic mechanism in eukaryotes that degrades and recycles long-lived cytoplasmic proteins, as well as damaged organelles. The hallmark of autophagy is the formation of double-membrane vesicles known as autophagosomes. I provided the initial evidence that CVB3 infection induces the formation of autophagosomes. Up-regulation of autophagosome formation enhances CVB3 replication, whereas downregulation of autophagy pathway reduces CVB3 replication. My results help clarify the nature of the intracellular membranes previously shown to be required for viral replication. For project 2, I demonstrated that CVB3 manipulates SRF expression via protein cleavage. SRF is a transcription factor vital for the expression of cardiac contractile/regulator genes, as well as gene silencing microRNAs. Cardiac-specific knockout of SRF in adult transgenic mice results in disruption of cardiac gene expression and development of severe DCM. I showed that SRF is cleaved in CVB3-infected mouse hearts and cardiomyocytes. Further studies revealed that SRF is cleaved at the 327 amino acids position by CVB3-encoded protease 2A. I demonstrated that SRF cleavage contributes to DCM by abolishing the transactivation property of SRF and generating dominant-negative SRF-truncates. Taken together, these novel viral strategies bridged existing knowledge and may serve as therapeutic targets for viral myocarditis/DCM.  ii  Preface The portion of following dissertation includes chapters which are based on two manuscripts, in that Chapter 3 is based on an article published in the Journal of Virology [Wong J, Zhang J, Si X, Gao G, Mao I, McManus BM, and Luo H (2008). Autophagosome supports coxsackievirus B3 replication in host cells. J Virol., 82(18):9143-9153], and Chapter 4 is based on another article published in Cell Research [Wong J, Zhang J, Yanagawa B, Luo Z, Yang X, Chang J, McManus B, and Luo H (2012). Cleavage of serum response factor mediated by enteroviral protease 2A contributes to impaired cardiac function. Cell Res., 22:360-371]. Specifically, I contributed to all experiments and analysis in Chapter 3 and Chapter 4, except in vivo coxsackievirus infection and heart function measurement, as well as cardiac gene expression measurement by microarray and subsequent validation by quantitative real-time PCR. In addition, parts of the contents from my review article published in Cardiovascular Research [Luo H, Wong J, and Wong B (2010). Protein degradation systems in viral myocarditis leading to dilated cardiomyopathy. Cardiovasc Res. 85(2):347-356] and book chapter published in Myocarditis [Wong J and Luo H (2011). Impaired cardiac function in viral myocarditis, Myocarditis, Cihakova D (Ed.), ISBN: 978-953-307-289-0, InTech, Rijeka, Croatia] are also used in this dissertation. During my PhD studies, I also conducted additional projects that resulted in two other firstauthor manuscripts: Wong J, Zhang J, Si X, Gao G, Luo H (2007). Inhibition of the extracellular signalregulated kinase signaling pathway is correlated with proteasome inhibitor suppression of coxsackievirus replication. Biochem Biophys Res Commun., 358(3):903-907., and Wong J, Si X, Zhang J, Angeles A, Luo H (2012). Degradation of adenosine-uridine binding factor 1: a strategy for coxsackieviral replication. Manuscript in preparation. In addition, I was involved in a number of related projects that resulted in eight co-author peer reviewed articles. This study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the Canadian Council on Animal Care. The protocol was approved by the Committee on Animal Care of the University of British Columbia (A03-0233).  iii  Table of contents Abstract ......................................................................................................................................................... ii Preface ......................................................................................................................................................... iii Table of contents..........................................................................................................................................iv List of tables ................................................................................................................................................ vii List of figures .............................................................................................................................................. viii List of abbreviations ..................................................................................................................................... ix Acknowledgements..................................................................................................................................... xii Chapter 1 – Introduction............................................................................................................................... 1 1.1.  Overview of myocarditis ............................................................................................................... 1  1.2.  Coxsackievirus ............................................................................................................................... 4  1.2.1  Discovery of coxsackievirus................................................................................................... 4  1.2.2  Classification and Properties of coxsackievirus .................................................................... 4  1.2.3  Lifecycle of coxsackievirus B3 ............................................................................................... 5  1.2.4  Functions of coxsackievirus-encoded proteins ..................................................................... 7  1.3.  Pathogenesis of CVB3-induced myocarditis: a tri-phasic disease................................................. 8  1.3.1  Viremic phase........................................................................................................................ 8  1.3.2  Inflammatory phase .............................................................................................................. 8  1.3.3  Remodeling phase................................................................................................................. 9  1.4.  Diagnosis and treatment of viral myocarditis ............................................................................. 11  1.5.  Research hypothesis and objective............................................................................................. 12  Chapter 2 – Materials and methods ........................................................................................................... 14 2.1  Materials ..................................................................................................................................... 14  2.2  Cell culture .................................................................................................................................. 14  2.3  In vitro and In vivo CVB3 infection .............................................................................................. 14  2.4  Heart function measurements .................................................................................................... 15  2.5  Plaque assay ................................................................................................................................ 15  2.6  Western blot analysis .................................................................................................................. 16  2.7  Immunocytochemistry ................................................................................................................ 16  2.8  Transmission electron microscopy ............................................................................................. 16  2.9  Microscopic monitoring of autophagosome formation ............................................................. 17  iv  2.10  RNA interference ........................................................................................................................ 17  2.11  Generation of SRF mutant constructs......................................................................................... 17  2.12  Luciferase assay .......................................................................................................................... 17  2.13  Affymetrix array analysis............................................................................................................. 18  2.14  Real-time quantitative RT-PCR .................................................................................................... 18  2.15  Statistical analysis ....................................................................................................................... 18  Chapter 3 – Autophagy in coxsackievirus infection .................................................................................... 19 3.1  Background ................................................................................................................................. 19  3.1.1  Autophagy ........................................................................................................................... 19  3.1.2  Autophagy in cardiac diseases ............................................................................................ 24  3.1.3  Autophagy in viral infections .............................................................................................. 24  3.2  Rationale ..................................................................................................................................... 25  3.3  Hypothesis and specific aims ...................................................................................................... 25  3.4  Results ......................................................................................................................................... 26  3.4.1  CVB3 infection increases autophagosome formation. ....................................................... 26  3.4.2  Mechanism of induction of autophagosome formation..................................................... 30  3.4.3  Autophagy inhibitor 3-MA reduces CVB3 replication. ........................................................ 31  3.4.4  Induction of autophagy enhances viral growth. ................................................................. 33  3.4.5  Knockdown of the genes critical for autophagosome formation reduces CVB replication.35  3.4.6  Expression of p62 following CVB3 infection. ...................................................................... 38  3.4.7  Lysosomal inhibition enhances CVB3 replication. .............................................................. 39  3.5  Discussion.................................................................................................................................... 41  3.6  Limitations and solutions ............................................................................................................ 43  3.7  Future directions ......................................................................................................................... 45  3.8  Summary ..................................................................................................................................... 46  Chapter 4 – Serum response factor in CVB3-induced myocarditis ............................................................. 47 4.1  Background ................................................................................................................................. 47  4.1.1  Mechanisms of CVB3-induced DCM ................................................................................... 47  4.1.2  Serum response factor ........................................................................................................ 49  4.1.3  SRF-mediated cardiac gene expression .............................................................................. 51  4.1.4  SRF in heart diseases ........................................................................................................... 52  4.2  Rationale ..................................................................................................................................... 53 v  4.3  Hypothesis and specific aims ...................................................................................................... 53  4.4  Results ......................................................................................................................................... 54  4.4.1  Cardiac function and cardiac gene expression following CVB3 infection ........................... 54  4.4.2  Expression levels of SRF following CVB3 infection.............................................................. 56  4.4.3  Cleavage of SRF following CVB3 infection .......................................................................... 59  4.4.4  Identification of SRF cleavage site during CVB3 infection .................................................. 63  4.4.5 Cellular localization and transcriptional function of CVB3-mediated SRF cleavage fragments ............................................................................................................................................ 67 4.4.6  Effects of CVB3-mediated SRF cleavage on viral replication .............................................. 70  4.4.7  Attenuation of SRF cleavage using peptide-based inhibitor............................................... 71  4.5  Discussion.................................................................................................................................... 73  4.6  Limitations and solutions ............................................................................................................ 75  4.7  Future directions ......................................................................................................................... 76  4.8  Summary ..................................................................................................................................... 77  Chapter 5 – Closing remarks ....................................................................................................................... 79 5.1  Research summary and conclusion............................................................................................. 79  5.2  Research significance .................................................................................................................. 80  Bibliography ................................................................................................................................................ 82  vi  List of tables Table 1. Primary etiological agents of myocarditis ....................................................................................... 3  vii  List of figures Figure 1. Lifecycle of coxsackievirus. ............................................................................................................ 6 Figure 2. Pathogenesis of viral myocarditis: a tri-phasic disease. .............................................................. 10 Figure 3. Process of autophagy pathway. ................................................................................................... 20 Figure 4. Molecular machineries of autophagy pathway. .......................................................................... 22 Figure 5. CVB3 infection increases autophagosome formation. ................................................................ 28 Figure 6. CVB3 infection induces eIF2α phosphorylation. .......................................................................... 30 Figure 7. Autophagy inhibitor 3-MA reduces CVB3 replication. ................................................................. 32 Figure 8. Induction of autophagy enhances viral replication. .................................................................... 34 Figure 9. Gene silencing of ATG7 by siRNA inhibits CVB3 replication. ....................................................... 36 Figure 10. Knockdown of class III PI3K components reduces CVB3 replication. ........................................ 37 Figure 11. Expression of p62 is not affected by CVB3 infection. ................................................................ 38 Figure 12. Lysosomal inhibition enhances CVB3 replication. ..................................................................... 40 Figure 13. Coxsackieviral protease 2A cleaves dystrophin at its 3' hinge................................................... 48 Figure 14. Serum response factor regulates cardiac gene expression. ...................................................... 50 Figure 15. Cardiac function and cardiac gene expression following CVB3 infection. ................................. 55 Figure 16. Protein and gene expression of SRF following CVB3 infection. ................................................. 57 Figure 17. Cleavage of SRF following CVB3 infection. ................................................................................ 60 Figure 18. Effect of caspase inhibition on CVB3-induced cleavage of SRF. ................................................ 62 Figure 19. Cleavage of SRF by overexpressing viral protease 2A................................................................ 64 Figure 20. Cleavage of SRF after T326 following CVB3 infection. ............................................................... 66 Figure 21. Cellular localization and transcriptional activity of SRF mutants. ............................................. 69 Figure 22. Effects of SRF dysregulation on CVB3 replication. ..................................................................... 70 Figure 23. Attenuation of SRF cleavage using peptide-based inhibitor...................................................... 72 Figure 24. Cleavage of SRF by viral protease 2A leads to myocyte dysfunction......................................... 78  viii  List of abbreviations 3-MA – 3-methyl adenine Ambra1 – Activating molecule in beclin-1-regulated autophagy ANP – Atrial natriuretic factor ATG – Autophagy related gene BSA – Bovine serum albumin CAR – Coxsackievirus and adenovirus Receptor CREB – cAMP responsive element binding protein CPE – cytopathic effect CVA – Coxsackievirus A CVB – Coxsackievirus B DAF – Decay accelerating factor DAPI – 4’,6-diamidino-2-phenylindole DCM – Dilated cardiomyopathy DMEM - Dulbecco's modified eagle medium eIF2α - Eukaryotic initiation factor 2α eIF4G – Eukaryotic initiation factor 4G ER – Endoplasmic reticulum EMB – Endomyocardial biopsy EMSA – Electrophoretic mobility shift assay FIP2000 – Focal adhesion kinase family interacting protein of 200 kD FMK – Fluoromethyl ketone GFP – Green fluorescent protein HEV – Human enterovirus IFN-γ – Interferon-γ  ix  IL – Interleukin IRES – Internal ribosome entry site LC3-PE – Microtubule-associated protein light-chain 3-phosphatidylethanolamine LVPW – Left ventricular posterior wall MADS – MCM1, Agamous, Deficiens, SRF MEF2 – Myocyte enhancer factor-2 MHC – Myosin heavy chain miRNA - MicroRNA MLC – Myosin light chain MOI – Multiplicity of infection mTOR – Mammalian target of rapamycin NFAT – Nuclear factor of activated T-cells NLS – Nuclear localization signal NK cells – Natural killer cells OCT-1 – Octamer-binding transcription factor ORF – Open reading frame PABP – Poly-A binding protein PBS – Phosphate buffered saline PCR – Polymerase chain reaction PERK – Protein kinase RNA-like endoplasmic reticulum kinase PFU – Plaque forming unit PI3K – Phosphoinositide-3 kinase PI3P – Phosphatidylinositol 3-phosphate PKR – Protein kinase R RT-PCR – Reverse-transcriptional PCR  x  SERCA – Sarcoplasmic reticulum Ca2+-ATPase SCID – Severe combined immunodeficiency SRE – Serum response element SRF – Serum response factor siRNA – Small interfering RNA ssRNA – Single-stranded RNA TBP – TATA-binding protein TBS – Tris-buffered saline TBST – TBS containing 0.1% Tween-20 TNF-α – Tumor necrosis factor-α ULK1/2 – Uncoordinated 51-like kinase 1/2 UTRs – Untranslated regions VP1 – Viral capsid protein-1 VPg-pU-pU – Uridylylated VPg Vps34 – Vacuolar protein sorting-34  xi  Acknowledgements I dedicate this thesis to my dear parents, Frank and Serena, who selflessly put my career success at their top priority. I cannot thank them enough for their love and care, and most importantly for their guidance in life. I would like to thank my long-time girlfriend and now my wife Cecilia for her loving, sharing, support, and understanding. She believes strongly in me and thinks highly of my ideas. She cooks me fish head soup to help me through “failed experiments” days. Cecilia is my muse. I attribute my fundamentalist thinking to her seemingly-random purchase of self-help books and her holistic nutrition schooling. I am enormously thankful for my supervisor Dr. Honglin Luo. She provided a harmonious environment for my post-undergraduate journey. Her excellence in supervision and mentorship inspires me to want the same for my trainees. She balances research guidance with encouraging independent thinking. I thank her trust in me and gave me freedom and resources to explore my research. I also thank her in giving me opportunities to take part in various projects and to interview and supervise undergraduate students. I also like to thank my co-supervisor Dr. Bruce McManus for his vision, leadership, and steadfast encouragement. I would also like to acknowledge my colleagues at the James Hogg Research Centre. Our humble technician Jingchun Zhang not only gave me great help in completing my projects but also shared his knowledge of traditional Chinese medicine and history with me. Post-doctoral fellow Dr. Xiaoning Si showed me the excitement and possibility of scientific experiments. I would like to thank Dr. Mary Zhang for her scientific guidance. My lab buddies David Lin, Guang Gao, Junyan Shi, Gabrial Fung, Tak Poon, Archie Angeles, Chris Hwang, Jayant Shravah, Allen Xiao, and Ivy Mao that make Luo lab the fun place to hang out. Special thanks to Dr. David Walker and Fanny Chu for guiding me through my electron microscopy work. I would like to thank all my committee chairs (Drs. Colin Fyfe and Haydn Pritchard) and members (Drs. Decheng Yang and Jian Ye) for their commitments to my many progress meetings and their discussions and suggestions on my research. The project presented in this thesis was supported by the Heart and Stroke Foundation of British Columbia and Yukon Grant-In-Aid (2008-2011) and Canadian Institute of Health Research (CIHR) Grant (2009-2012).  xii  My graduate study was supported by the CIHR Canadian Graduate Scholarship (Master Award, 2007-2009), Michael Smith Foundation of Health Research Junior Graduate Studentship (2007-2009), and Frederick Banting and Charles Best Canada Graduate Scholarship (Doctoral Award, 2009-2012).  xiii  Chapter 1 – Introduction 1.1.  Overview of myocarditis Myocarditis is defined clinically as inflammation of the myocardium and necrosis and/or  degeneration of adjacent myocytes in the absence of an ischemic event (9, 137). It is a major cause of sudden, unexpected death in individuals less than 40 years old (54). More importantly, it is a potential precursor to dilated cardiomyopathy (DCM), which leads to congestive heart failure (36, 210). Each year ~100,000 new cases of DCM are diagnosed in the United States alone and the 10-year survival rate of the disease is ~40% (202, 204). Unfortunately, there is no specific and effective therapy available to date for the treatment of myocarditis/DCM. Currently, supportive therapies are primarily used for myocarditis/DCM patient care. At end-stage DCM, heart transplantation represents the only definitive therapeutic option (138). According to the International Society of Heart and Lung Transplantation Registry, cardiomyopathy accounts for about half of all patients requiring heart transplants (87). Myocarditis is originally described by Joseph Sobernheim in 1837 as inflammation of the myocardium (164). Remarkable advances in our understanding, diagnosis, and treatment of myocarditis were made in the last 150 years. Myocarditis is an insidious disease that is typically asymptomatic and subclinical, thus conceals its true incidence. It was post-mortem studies that revealed the astounding prevalence of myocarditis (54). The advent of transvenous endomyocardial biopsy in 1962 by Sakakibara and Konno opened up opportunity to assess the premorbid presence of myocardial inflammation in patients (173). Yet, different biopsy studies derived highly variable incidences of myocarditis, ranging from 0 to 80 percent (122). The disparities were attributed to differences in patient selections, diagnostic criteria, as well as the inherent insensitivity of the biopsy in the detection of inflammatory disease (38, 104, 174). To resolve the disparities and standardize diagnostic evaluation, the Dallas criteria for the histological diagnosis of myocarditis were introduced in 1986 (8). These criteria include categories of acute (lymphocytic infiltration with myocytolysis), borderline (lymphocytic infiltration without myocytolysis), and persistent (repeatedly observed lymphocytic infiltration and myocytolysis on serial biopsies), as well as negative (absent of lymphocytic infiltration and myocytolysis) (8). However, the Dallas criteria probably still underestimates the true incidence of myocarditis. The causes of myocarditis are diverse (summarized in Table 1) and include infectious (viral, bacterial, parasitic, and fungal) agents, toxins, allergens, and immunologic disorders (61, 70, 139). Infectious agents are the most common cause of myocarditis worldwide. The relevance of specific infectious etiologies varies with geographical regions. In Central and South America, myocarditis is  commonly associated with Chagas disease caused by the parasite Trypanosoma cruzi, whereas in North America and other developed nations, cardiotropic viruses are the primary infectious agents of myocarditis (5, 151, 204). Routine serological examinations of serum from patients during acute myocarditis and convalescence demonstrated rising antibody titers against selected viruses (27). Furthermore, viral genome has been identified in the myocardium of patients with clinically suspected myocarditis and DCM. Endomyocardial biopsy examinations of myocarditis and DCM patients by polymerase chain reaction (PCR) revealed the presence of viral genomes in one-third of the inspected cases (94, 177). The presence of viral genome is associated with poor prognosis. The two-year mortality rate in patients tested positive for viral genome in their endomyocardial biopsies is nine times higher than those tested negative (202). Taken together, these studies implicated a causal link between viral myocarditis and the development of DCM. More than twenty common viruses, including enteroviruses, adenovirus, parvovirus, cytomegalovirus, hepatitis C virus, influenza virus, and human immunodeficiency virus, have been associated with myocarditis in humans (77). Among them, enteroviruses have been the most widely studied viruses in the pathogenesis of viral myocarditis. Global surveillance data compiled by the World Health Organization indicate that coxsackievirus of the enterovirus genus is the dominant cause of myocarditis (77). Young children, particularly newborns under 6 months of ages, are extremely susceptible to coxsackievirus infections. Myocarditis is responsible for approximately 20% of sudden unexpected deaths among young children (54, 163). Thus far, no effective vaccine is available for coxsackievirus infection. The only definitive treatment currently available for DCM is heart transplantation.  2  Table 1. Primary etiological agents of myocarditis Viruses  Bacteria  Parasites  Fungi  • Adenovirus • Coxsackievirus • Cytomegalovirus • Epstein-Barr virus • Hepatitis C virus • Human immunodeficiency virus • Influenza virus • Parvovirus B19  • Chlamydia pneumonia • Corynebacterium diphtheriae • Mycoplasma pneumoniae • Serratia marcescens • Streptococcus pneumoniae • Treponema pallidum  • Echinococcus granulosus • Taenia solium • Toxoplasma gondii • Trypanosoma cruzi • Visceral larva migrans • Wuchereria bancrofti  • • • • • • • • •  Toxins  Allergens  Immunologic disorders  • • • • • • • •  • • • • • • • •  • • • • • • • • • •  Amphetamines Anthracyclines Cocaine Catecholamines Ethanol Fluorouracil Heavy metals Interleukin-2  Acetazolamide Amitriptyline Cefaclor Colchicine Lidocaine Penicillin Tetanus toxoid Tetracycline  Actinomyces Aspergillus Blastomyces Candida Coccidioides Crytoccoccus Histoplasma Mucormycoses Nocardia  Churg-Strauss syndrome Inflammatory bowel disease Insulin-dependent diabetes mellitus Myasthenia gravis Sarcoidosis Systemic lupus erythematosus Takayasu’s arteritis Thyrotoxicosis Kawasaki’s disease Wegener’s granulomatosis  Multiple etiologies have been identified for the myocarditis, but cardiotropic viruses are the most common causes of the disease in North America and other developed nations. Among them, coxsackievirus of the enterovirus genus is the dominant cause of myocarditis. Data are from Feldman and McNamara (61).  3  1.2.  Coxsackievirus  1.2.1  Discovery of coxsackievirus Coxsackievirus was originally isolated by Gilbert Dalldorf and Grace Sickles in 1948 while  searching for a cure for polio (45). Previous work by Dalldorf suggested that fluid collected from monkeys infected with a non-poliovirus preparation have protective effect against polio. Dalldorf and Sickles then attempted to isolate such polio-protective viruses using newborn mice and fecal specimens from two young boys suffering from flaccid paralysis (45). These non-polio isolates differed from poliovirus that they were able to infect newborn (suckling) mice in addition to primates. Infection by these non-polio isolates often mimicked mild or non-paralytic poliovirus infection. The newly discovered virus family was eventually given the name Coxsackie, for the Hudson River town of Coxsackie, New York, where Dalldorf obtained the initial fecal specimens. In the following year, Dalldorf and Sickles reported a virus, later called coxsackievirus A (CVA), which induced fatal paralysis and myositis in suckling mice, but not adult mice (43). In the year after, Edward Curnen, Ernest Shaw, and Joseph Melnick discovered the coxsackievirus B (CVB) that confers different antigenicity and pathogenicity from CVA (142). Suckling mice infected with CVB resulted in focal and limited myositis in striated muscles, as well as degeneration of brain, pancreas, heart, and embryonic fat pads (141). 1.2.2  Classification and Properties of coxsackievirus Coxsackievirus, a family member of Picornaviridae, belongs to the human enterovirus (HEV)  genus. There are twenty-three known serotypes (1-22, 24) of CVA and six serotypes (1-6) of CVB, which belong to HEV species A (CVA2-8, 10, 12, 14, and16), B (CVB1-6, CVA9 and 23), or C (CVA1, 11, 13, 17, 19-22, and 24) (199). CVA serotypes mainly cause enteric diseases, while the six CVB serotypes are associated with more severe diseases such as aseptic meningitis, pancreatitis, and myocarditis (44). Among the CVBs, only CVB1, 3, and 5 are cardiotropic (72). Coxsackievirus B3 (CVB3) is a non-enveloped single-positive-stranded RNA virus. The CVB3 genome is a single-stranded sense RNA (ssRNA(+)) molecule of ~7400 nucleotides in length. It comprises a single open reading frame (ORF) flanked on both ends by untranslated regions (UTRs). The ORF encodes 11 viral proteins that are translated into a large polyprotein of 2185 amino acids, including four structural proteins (VP1, VP2, VP3, and VP4), two viral proteases (2A and 3C), an RNA-dependent-RNApolymerase (3D), a small polypeptide that serve as primer for RNA synthesis initiation (3B, also known as, VPg), an ATPase (2C), two host membrane modifiers (2B and 3A). In addition, there are three functional intermediates (2BC, 3AB, and 3CD) (89). The 5’UTR contains ~800 bases that folds into highly  4  ordered secondary structures, including the internal ribosome entry site (IRES) that directs viral genome translation initiation. Unlike eukaryotic mRNA, the 5’UTR of viral genome is covalently linked to a small viral polypeptide VPg, instead of the 7-methylguanosine triphosphate cap (63). These structural features of the viral genome 5’UTR serve an important function in cap-independent translation that plays a role in subverting host protein translation machineries for viral replication as discussed in the later sections. On the other hand, the 3’UTR (~100 bases) of viral genome is polyadenylated. The CVB3 genome is encapsulated in an icosahedral protein capsid of ~30 nm in diameter (172). The capsid is composed of 4 viral structural proteins: VP1, VP2, VP3, and VP4. One molecule of each structural protein combines and forms a protomer. Five protomers in turn compose a pentamer and 12 pentamers assemble together to form the capsid. 1.2.3  Lifecycle of coxsackievirus B3 Coxsackievirus lifecycle (summarized in Figure 1) begins with its binding and clustering of co-  receptor DAF (decay accelerating factor, also known as CD55), which leads CVB3 to its main receptor CAR (coxsackievirus and adenovirus receptor) located in the intercalated disk of cardiomyocyte (179). Its engagement with CAR allows CVB3 to internalize into endosome through the clathrin-dependent endocytosis pathway (39). Subsequent acidification of endosome leads to viral uncoating and release of viral genome (ssRNA(+)) into the cytoplasm. Now, the viral genome translates into a large viral polyprotein, which is processed by viral proteases 2A and 3C embedded within, into 11 viral structural and non-structural proteins, as described previously (172). The RNA-dependent-RNA-polymerase 3D together with primer uridylylated VPg (VPg-pU-pU) transcribe ssRNA(+) into a viral RNA intermediate (ssRNA(-)) that in turn serves as template for the synthesis of multiple viral progeny genomes (ssRNA(+))(150, 152, 168). The viral progeny genomes and structural proteins are integrated to make complete virions. The lifecycle of CVB3 is completed by releasing viral progenies for new rounds of infections.  5  6 Progeny Release  CVB3 Lifecycle 1 Attachment  4 Replication  Internalization 2  Translation 3 Assembly 5  Viral RNA Viral Polymerase Viral Poly-protein Structural Protein Non-structural Protein  CAR DAF Endosome Membrane Platform  Figure 1. Lifecycle of coxsackievirus. Coxsackievirus lifecycle begins with its binding and clustering of co-receptor DAF (decay accelerating factor, also known as CD55), which leads CVB3 to its main receptor CAR (coxsackievirus and adenovirus receptor) located in the intercalated disk of cardiomyocyte. Its engagement with CAR allows CVB3 to internalize into endosome. Now, CVB3 uncoats and injects its viral genome (ssRNA(+)) into the cytoplasm. The viral genome translates into a large viral polyprotein, which is processed by viral proteases embedded within, into structural and non-structural proteins. The RNA-dependent-RNApolymerase 3D transcribe ssRNA(+) into a viral RNA intermediate (ssRNA(-)) that in turn serves as template for the synthesis of multiple viral progeny genomes (ssRNA(+)). The viral progeny genomes and structural proteins are integrated to make complete virions. The lifecycle of CVB3 is completed by releasing viral progenies for infections of nearby cells.  6  1.2.4  Functions of coxsackievirus-encoded proteins CVB3, like other family members of Picornaviridae, encodes two viral proteases 2A and 3C, as  well as intermediate 3CD. Both 2A and 3C are cysteine proteases and are homologous to trypsin-like family of serine proteases. Not only are these viral proteases important in the processing of viral polyprotein (82, 84), but they also contribute to successful viral replication by cleaving multiple host proteins involved in host RNA transcription, host protein translation, cell signalling, and cell structure. Protease 2A cleaves eukaryotic initiation factor 4G (eIF4G) and poly-A binding protein (PABP) (95, 100, 114, 115). Both eIF4G and PABP are essential for host cap-dependent translation. As a result, cleavage of both proteins halts cellular protein synthesis (100, 115). CVB3 protein synthesis, however, is not affected due to the uncapped nature of viral RNA and its cap-independent translation mechanism conferred by the internal ribosome entry site (IRES) located at the 5’UTR of viral genome (92). This viral strategy, thereby, efficiently subverts host protein synthesis machineries to focus on viral protein synthesis. Furthermore, viral protease 2A cleaves the cytoskeletal proteins dystrophin and cytokeratin-8 (11-13) to facilitate viral progeny release. Viral protease 3C, on the other hand, supports viral replication by shutting down host gene transcription via cleavage of cAMP response element binding protein (CREB), octamer-binding transcription factor (Oct-1), and cellular TATA-binding protein (TBP), thus adds cellular stress on infected cardiomyocytes (40, 206, 207). Both viral proteases also contribute to myocyte death by activating host apoptosis by cleaving Bcl-2 family protein Bid and inducing caspase-8-mediated activation of caspase-3 (16, 33, 74). In vitro observation of CVB3-infected cells reveals numerous degenerative cellular changes associated with the aforementioned pro-viral events, i.e. host transcription and translation shutdown and disruption of cytoskeleton, during CVB3 replication, collectively referred as cytopathic effect (CPE) (29, 30). Infected cells suffering from CPE undergo morphological changes such as cell rounding and membrane permeability alteration, followed by detachment from cell culture surface, and eventual cell lysis through apoptosis, necrosis, or both. Non-structural viral proteins other than the polymerase and proteases, such as 2B, 3A, and intermediate 2BC, also play important roles in CVB3 lifecycle. Experimental evidence suggests that viral 2B, 2BC, and 3A initiate intracellular membrane reorganization to support viral replication and increase plasma membrane permeability to facilitate viral progeny release (52, 194). This is supported by the observation that poliovirus carrying a 2B mutation is defective in RNA amplification and viral progeny release in cell culture (96).  7  1.3.  Pathogenesis of CVB3-induced myocarditis: a tri-phasic disease Coxsackievirus B3 infection may lead to severe myocardial injury. The outcome of CVB3  infection depends on the complex interaction between pro-viral mechanisms and host anti-viral immune responses (37, 126, 138). Data from murine experimental model of CVB3-induced myocarditis suggests that the disease progression can be characterized in three distinct phases: (I) viremic, (II) inflammatory infiltration, and (III) cardiac remodeling (125, 189). Featured events in each phase of viral myocarditis are summarized in Figure 2. 1.3.1  Viremic phase The viremic phase (0-4 days post infection) features prominent viral replication and early  cardiomyocyte damages without significant host immune responses during acute myocarditis (125, 189). It is well documented that CVB3 inflicts direct damages to infected cardiomyocytes, which contributes significantly to subsequent disease development (58, 127). As early as 2 days post infection, cytopathic lesions featuring single or multi-cell vacuolar changes are observed in CVB3-infected immune competent mice. Numerous and widespread islands of cardiomyocytes with cytopathic lesions, contraction band coagulation necrosis, as well as early single-cell calcification are observed from day 3 onwards (37, 138). Furthermore, in situ hybridization analysis shows co-localization of viral genome with damaged cardiomyocytes (105, 127). Further evidence of direct virus-induced myocardial injury comes from CVB3 infection of SCID (severe combined immunodeficiency) mice, which has no mature T and B cells due to their inability to complete V(D)J gene recombination. CVB3-infected SCID mice develop an early and extensive cardiomyocyte coagulative necrosis and contraction band necrosis as compared to their wildtype counterparts (37). 1.3.2  Inflammatory phase The inflammatory infiltration phase (5-14 days post infection) takes place in response to viremia  (125, 189). Natural killer (NK) cells and macrophages detect viral invasion and evoke the innate immune response, resulting in inflammatory cell infiltration of the myocardium. The macrophages and NK cells drive viral clearance by mediating cytolysis of infected cells, and by producing pro-inflammatory cytokines including tumour necrosis factor-α (TNF-α), interleukin-1 (IL-1), IL-2, and interferon-γ (IFN-γ) (58, 127). The innate immune response also mounts an adaptive immune response executed by antigenspecific T-lymphocytes (CD8+ cytotoxic T-cells and CD4+ T-helper cells) and antibody-producing Blymphocytes, which help eliminate viral particles and viral-infected cells (58). However, these well-  8  intentioned immunologic host defense mechanisms, if persistently activated, become detrimental and exacerbate myocardial injury. Indeed, certain pro-inflammatory cytokines augmented upon CVB3 infection, such as TNF-α and IL-1, are shown to promote CVB3-induced myocarditis in otherwise disease resistant mice (116). Furthermore, specific removal of certain immune cell populations such as Tlymphocytes in experimental myocarditis mouse models proves to be beneficial (165). These mice have markedly reduced inflammatory infiltrates and mortality rate upon CVB3 challenge. The humoral and cellular immune responses become pathologic when they morph into self-attacking entities upon exposure to secluded cardiac-antigens released from damaged cardiomyocytes, such as cardiac myosin, troponin-1, and vimentin (110, 127). Auto-antibodies against these cardiac proteins were detected in the sera of patients with myocarditis and DCM (47). Some of these auto-antibodies are also shown to cross-react with CVB3 particles (131), supporting the “antigenic mimicry” hypothesis in viral myocarditis (71). All in all, these pathologic immune reactions further contribute to myocyte injuries in the subacute and chronic stages of the disease, leading to heart failure and death. 1.3.3  Remodeling phase The cardiac remodelling phase (15 days post infection onwards) serves to repair the damaged  myocardium in the absence or presence of low-level persistent viral genomes (125, 189). Due to the terminally differentiated nature of cardiomyocytes, myocytes loss in the prior phases of viral myocarditis cannot be replaced through cell proliferation. In addition, viral genomes persisted in the forms of replication-defective mutants and double-stranded RNAs (as well as their consequential sporadic viral protein production) may exacerbate myocyte loss in the long term by chronically activate the immune response in the myocardium (62). Progressive fibrotic reparation takes place in regions of myocyte loss, resulting in abundant collagen accumulation in between remaining viable myocytes. The extensive fibrotic scars, however, are ineffective in facilitating contractile force transmission and electrical signal transduction. Still, cardiac output must be maintained. Efforts of the imperfect “band-aids” and remaining myocytes in delivering adequate cardiac output drive the continual restructuring of the myocardium, with deterioration and further reparation of the interstitial collagen network and cardiomyocyte hypertrophy. However, cardiac output cannot be sustained in the long term. Over time, progressive decompensatory molecular and structural changes take place in the myocardium. Ultimately, this results in myocardial wall thinning and left ventricle dilatation, leading to DCM and heart failure (64, 135).  9  Phase I Viremic 0 Viremia Acute myocyte damage Focal microcalcification Macrophage activation Cytokine expression (IL-1, IL-2, TNF-α α , IFN-γγ)  Phase II Inflammatory 4  Phase III Cardiac remodeling 14  Virus clearing Natural Killer cells T cells & B cells infiltration Neutralizing antibodies  90 days  Virus absence Viral RNA persistence Myocardial fibrosis Dilated cardiomyopathy Heart failure  Figure 2. Pathogenesis of viral myocarditis: a tri-phasic disease. Pathogenesis of viral myocarditis: a tri-phasic disease. In experimental mouse model, three distinct phases of viral myocarditis has been characterized: viremic, inflammatory, and remodeling phases. The viremic phase features prominent viral replication with acute myocyte damage and focal microcalcification in the absence of inflammatory infiltrates. Initial release of cytokines such as IL-1, IL-2, TNF-α, and IFN-γ in infected heart attracted macrophages and NK cells to the sites of injury, which initiates the inflammatory phase characterized by the infiltration of T cells and B cells, as well as the production of neutralizing antibodies for viral clearance. In the remodeling phase of myocarditis, viral particles are typically absent in blood and peripheral tissues. Persistence of viral genome and immune infiltrates may contribute to long-term tissue degeneration and progression of viral myocarditis to DCM and heart failure. This figure is modified from Feldman and McNamara (61).  10  1.4.  Diagnosis and treatment of viral myocarditis Viral myocarditis is a presumptive diagnosis based on history and clinical features. Viral  myocarditis patients may present with a history of recent flu-like symptoms of fever, arthralgia, and malaise. Clinical presentations of viral myocarditis varied widely. Similar to other forms of myocarditis, viral myocarditis is typically asymptomatic and subclinical. However, asymptomatic patients may have electrocardiographic abnormalities. Severe patients with viral myocarditis may have signs and symptoms of clinical heart failure (e.g. shortness of breath, chest pain, and ventricular arrhythmia) and ventricular dilatation. Patients with fulminant viral myocarditis may have severe left ventricular dysfunction, with or without cardiac dilatation (48). Clinical laboratory testing may reveal elevations in leukocyte count, sedimentation rate, and cardiac fraction of creatine kinase, troponin T, as well as troponin I ((49, 118, 181, 195)). Elevated serum levels of cardiac creatine kinase and troponins are indicative of myocardial cell damage. Electrocardiogram may show ventricular arrhythmias. Serological examination may also reveal circulation of anti-viral antibodies (55, 119, 120). Endomyocardial biopsy (EMB) assessed by the Dallas criteria is still the reference standard for diagnosing viral myocarditis, despite its limited sensitivity and specificity (18, 24, 55). Contrast-enhanced magnetic resonance imaging and echocardiography may help localizing and assessing the extent of inflammation in patients with presumed myocarditis (66, 123). Follow-up EMB sampling at the abnormal myocardial area may enhance its diagnostic value. Molecular techniques such as reverse-transcriptional PCR (RT-PCR) and in situ hybridization allow detection of the presence of viral genome in EMB specimens (51, 130, 204). Akin to other forms of myocarditis, the treatment of viral myocarditis is mainly designed around supportive measures, such as inotropic therapy, afterload reduction, arrhythmia management, and adequate tissue oxygenation (61, 192). Although the activation of cellular and humoral autoimmunity may play a role in the long-term progression of myocarditis, immunosuppressive therapy is not recommended in patients with viral myocarditis (1). Various randomized, placebo-controlled trials have failed to demonstrate beneficial effects of immunosuppression (117, 140, 167). Treatment with specific antiviral drugs for viral myocarditis such as Isoxazoles (WIN54954) and Pleconaril (VP63843) has been shown to be beneficial (65, 132). These inhibitors are broad-spectrum viral capsid function inhibitors that block enteroviral attachment, entry, and uncoating. A clinical trial involving Pleconaril usage in children and adults patients with enteroviral meningitis and upper respiratory tract infections were reported to decrease severity and duration of the diseases (19). Still, effective therapeutics for the treatment of viral myocarditis/DCM is greatly needed to improve overall patient survival and cardiac function.  11  1.5.  Research hypothesis and objective Viral myocarditis, the inflammation of myocardium initiated by viral infection, is an important  cause of mortality in newborns and neonates. In addition, it is a precursor to dilated cardiomyopathy. To date, no effective therapy is available for viral myocarditis/DCM. Coxsackievirus B3 is a primary human pathogen of viral myocarditis. Extensive research efforts on CVB3 have broadened our understanding of viral lifecycle. However, the pathogenesis of coxsackievirus-induced myocarditis is not fully understood. The objective of this dissertation is to explore the role of host protein manipulation in coxsackieviral replication and pathogenicity. My hypotheses are (1) coxsackievirus hijacks host’s cellular autophagy mechanism to facilitate its own replication; and (2) the serum response factor (SRF) is cleaved by viral protease 2A during coxsackievirus infection and contributes to impaired myocardial function and progression to DCM. As described previously, the general gist of viral lifecycle includes viral particle internalization upon receptor engagement, followed by cytosolic release of viral genomic RNA and viral protein synthesis. Viral proteins manipulate multiple host pathways to facilitate the amplification of viral genomes and viral proteins. Viral proteins 2B, 2BC, and 3A induce massive intracellular membrane reorganization, resulting in vacuolization of host cells. These viral-induced membrane structures are said to support viral replication by serving as physical scaffolds for the assembly of viral replication complexes. However, the nature of these membranes remains unknown. In Chapter 3, I explored whether CVB3 manipulates the host autophagy protein degradation pathway to supply membrane platforms for successful viral replication. Aside from detail events in viral replication, the underlying molecular mechanism that impairs heart function and drives the progression of viral myocarditis and DCM is also vaguely understood. As introduced earlier, the virus-encoded proteases play vital roles in CVB3 lifecycle. Not only are these viral proteases important in the processing of viral polyprotein, but they also contribute to successful viral replication by cleaving multiple host proteins involved in host RNA transcription, host protein translation, cell signalling, and cell structure. Viral proteases have been recognized as important pathogenic factors contributing to the development of DCM. It has been shown that transgenic mice with inducible, cardiac-specific expression of viral protease 2A develop severe cardiac dysfunction and DCM (205). However, the mechanism underlying viral protease-induced DCM is not fully understood. In Chapter 4, I explored whether coxsackieviral protease manipulates the expression of serum response factor, an essential transcription factor for the expression of cardiac contractile and regulator genes, and contributes to impaired heart function and progression of viral myocarditis and DCM.  12  Bridging the existing findings helps create a complete picture of CVB3-induced myocarditis that is valuable to the development of novel therapeutics to attenuate viral replication and ameliorate viralinduced injury. Combinations of antiviral therapies may improve efficacy while minimizing side-effects.  13  Chapter 2 – Materials and methods 2.1  Materials Rapamycin, ammonium chloride (NH4Cl), bafilomycin A, 3-methyladenine (3-MA), monoclonal  anti-β-actin antibody, and monoclonal anti-GAPDH antibody were purchased from Sigma. A monoclonal anti-VP1 antibody was obtained from DakoCytomation. Beclin-1 small interfering RNA (siRNA), lysosome-associated membrane protein 2 (LAMP2) siRNA, SRF siRNA, control scramble siRNA, polyclonal anti-SRF C-terminus antibody, anti-FLAG antibody, anti-eIF4γ antibody, and horseradish peroxidaseconjugated secondary antibodies were purchased from Santa Cruz Biotechnology. Polyclonal anti-SRF Nterminus antibody was from Novus Biological. A monoclonal anti-Vps34 antibody was obtained from Life Technologies. Vps34 siRNA, ATG7 siRNA, and scramble control siRNA were obtained from Qiagen. A monoclonal anti-Beclin-1 antibody was purchased from BD Biosciences. Polyclonal guinea pig anti-p62 was purchased from Progen. A polyclonal rabbit anti-LC3 antibody was obtained from MBL International. A polyclonal rabbit anti-phospho-eIF2α antibody was obtained from Cell Signaling. Polyclonal rabbit anti-ATG7 and anti-LAMP2 antibodies were obtained from Cell Signaling and Abcam, respectively. z-VAD-fmk was purchased from EMD Biosciences. z-LKST-fmk was custom synthesized by CPC Scientific.  2.2  Cell culture HeLa cells and HEK293A cells obtained from the American Type Culture Collection were used for  this study. HeLa cells were grown in complete medium (Dulbecco's modified Eagle's medium [DMEM] supplemented with 10% heat-inactivated newborn calf serum). HEK293A cells were maintained in complete medium with the addition of 1% nonessential amino acids. The majority of the experiments in Chapter 3 were performed with HEK293A cells, and critical experiments were repeated using Hela cells. The HL-1 cell line, a murine cardiac muscle cell line established from an AT-1 mouse atrial cardiomyocyte tumor lineage, was a generous gift from Dr William C Claycomb (Louisiana State University Medical Center, USA). Cells were plated and maintained in Claycomb medium (JRH Biosciences) supplemented with 10% fetal bovine serum, 100 μg/ml penicillin-streptomycin, 0.1 mM norepinephrine in ascorbic acid, and 2 mM L-glutamine as described previously (129).  2.3  In vitro and In vivo CVB3 infection HeLa cells were infected at a multiplicity of infection (MOI) of 10 with CVB3 (Kandolf strain) or  sham infected with phosphate-buffered saline (PBS) for 1 h in serum-free DMEM. The cells were then  14  washed with PBS and cultured in complete medium for various times until they were harvested. HEK293A cells and HL-1 cardiomyocytes were infected in a similar fashion, but with a higher MOI of 100. For autophagy induction, cells were either treated with rapamycin for 3 h or starved in Hank's balanced salt solution (137 mM NaCl, 5.4 mM KCl, 0.8 mM MgSO4, 0.8 mM Na2HPO4, 0.4 mM KH2PO4, 4.2 mM NaHCO3, 20 mM HEPES, and 1.5 mM CaCl2, pH 7.4) for 2 h prior to virus infection. For inhibition experiments, cells were infected with CVB3 in the presence or absence of various inhibitors or vehicle (dimethyl sulfoxide) for 1 h, washed with PBS, and then incubated with fresh DMEM containing different concentrations of inhibitors unless otherwise specified in the figure legends. Murine coxsackievirus infection was performed as described (208). Briefly, male A/J mice at the age of ~5 weeks were either infected intraperitoneally with 105 plaque-forming unit (PFU) of CVB3 or sham infected with PBS for 3, 9, and 30 days. This study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the Canadian Council on Animal Care. The protocol was approved by the Animal Care Committee of the University of British Columbia (#A03-0233).  2.4  Heart function measurements Heart function was measured before infection (day 0) and on days 3, 9, and 30 after CVB3  infection using two-dimensional echocardiography (Sonos 5500, Philips) with a 12 MHz S-12 probe (Sonos 5000). Parasternal long and short axis images at the level of the mitral valve and papillary muscles were taken and these measurements were normalized to changes in body weight and then utilized for calculation of functional parameters. All measurements were averaged for five consecutive cardiac cycles.  2.5  Plaque assay The virus titer in the cell supernatant was determined by an agar overlay plaque assay  performed in triplicate, as described previously (6). In brief, the supernatant from CVB3-infected cells was serially diluted 10-fold and overlaid on a 90 to 95% confluent monolayer of HeLa cells in 6-well plates. After 1 h of incubation (5% CO2, 37°C), the HeLa cells were washed with PBS to remove nonbinding viruses, followed by overlay with 2 ml of warm complete medium containing 0.75% agar in each well. The cells were incubated at 37°C for 72 h and then fixed with Carnoy's fixative (75% ethanol-25% acetic acid) and stained with 1% crystal violet. Plaques were counted, and the viral titer was calculated as PFU per milliliter.  15  2.6  Western blot analysis Western blotting was performed as described previously (26, 128). After treatment, the cells  were incubated with lysis buffer (50 mM Tris-HCl, pH 7.2, 250 mM NaCl, 0.1% NP-40, 2 mM EDTA, 10% glycerol) containing protease inhibitor cocktail (Roche) on ice for 10 min. Cell lysates were then harvested by scraping, followed by brief sonication and centrifugation at 12,000 × g for 10 min at 4°C. The protein concentration was determined by the Bradford assay (Bio-Rad). Equal amounts of protein were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis and then transferred to nitrocellulose membranes (Amersham). The membranes were blocked for 1 h with 5% nonfat dry milk Tris-buffered saline solution containing 0.1% Tween 20 (TBST). The blots were then incubated for 1 h with the primary antibody prepared in 2.5% milk TBST, followed by three times washing with TBST. The membrane was then incubation for another hour with a HRP-conjugated secondary antibody, followed by washing with TBST. Immunoreactive bands were visualized by enhanced chemiluminescence (SuperSignal West Pico kit; Pierce) and documented by ChemiGenius 2 (Syngene).  2.7  Immunocytochemistry Immunocytochemical staining was performed as described previously (68). Cells grown on glass  slides were washed with PBS and fixed in 4% formaldehyde. Cells were then permeabilized with 0.1% Triton X-100 for 10 min, blocked with 5% bovine serum albumin (BSA) in Tris-buffered saline (TBS) for 30 min. After incubation with primary antibodies at 4 °C overnight and then with anti-rabbit AlexaFluor 594 or anti-mouse AlexaFluor 488 IgG at room temperature for 1 h, cells were washed with PBS and then mounted on slides using VectaShield medium containing 4’,6-diamidino-2-phenylindole (DAPI). Cells were imaged and analyzed using a Leica SP2 AOBS confocal fluorescence microscope.  2.8  Transmission electron microscopy For ultrastructural analysis, HeLa cells and HEK293A cells were sham infected or infected with  CVB3 at MOIs of 10 and 100, respectively, for 5 and 7 h. Briefly, cells were fixed in 2.5% glutaraldehyde (Polysciences Inc.) in PBS, pH 7.2, for 10 min at room temperature. Following three washes in PBS, cells were postfixed in 1% osmium tetroxide (Polysciences Inc.), 1% potassium ferrocyanide, 100 mM sodium cacodylate buffer, pH 7.4, for 1 h at room temperature. After several rinses in distilled water, cells were dehydrated in a graded series of acetone washes and embedded in Eponate 12 resin (Ted Pella Inc.). Sections (70 to 80 nm) were cut and viewed on a Tecnai 12 transmission electron microscope (FEI Inc.). Approximately 15 cells were counted, and for each cell the number of double-membrane vesicles was  16  examined, as previously described (4). Autophagosomes were defined as double-membrane vacuoles measuring 0.2 to 1.0 μm.  2.9  Microscopic monitoring of autophagosome formation For the detection of autophagosomes, cells were transiently transfected with GFP-LC3 plasmid  (a generous gift from Sharon A. Tooze at the Cancer Research UK London Research Institute, United Kingdom), using Lipofectamine 2000 (Life Technologies). After 24 h of transfection, cells were sham or CVB3-infected for 5 h, and the fluorescence of GFP-LC3 was observed under a Leica SP2 AOBS confocal fluorescence microscope. The number of cells with punctate GFP-LC3 localization relative to all green fluorescent protein (GFP)-positive cells was counted (a minimum of 200 GFP-positive cells were counted in total for each condition) and presented as a percentage, as previously described (88).  2.10 RNA interference HeLa cells were grown at 50% confluence were transiently transfected with various smallinterfering RNAs (siRNAs) at a concentration of 50 to 80 nM, using Lipofectamine 2000 or HiPerFect (Qiagen) according to the manufacturer's instructions. A scrambled siRNA was used as a negative control. The silencing efficiency was detected by western blot analyses using the respective antibodies. After 48 h, cells were infected with CVB3 as described above. Cell supernatants and lysates were collected 7 h post infection and analyzed for viral titer and protein expression by plaque assay and western blotting, respectively.  2.11 Generation of SRF mutant constructs The pCMV-3×FLAG-SRF construct, a generous gift from Dr Ron Prywes (Columbia University, USA), was used as the template for the generation of site-directed mutagenesis (SRFG349E and SRFG327E mutants). Truncated SRF cDNAs were generated by PCR using the following primers: SRF-N forward 5’-GTCGACTCTAGAATGTTACCG-3’ and reverse 5’-AGGAGACGGATCCGCTTCATG-3’; SRF-C forward 5’-CTGAAGTCTAGAGGCAGCGG-3’ and reverse 5’-ACCCGGGATCCTTTAGATCAT-3’. SRF-N and SRF-C cDNAs were subcloned into pCMV-3×FLAG vector using XbaI and BamHI restriction digestions.  2.12 Luciferase assay HeLa and HL-1 cells at 90% confluence in 24-well tissue culture plates were transiently cotransfected (using Lipofectamine 2000) with a Firefly luciferase reporter construct containing cardiac αactin promoter (32) and a Renilla luciferease control vector (Promega) together with plasmids encoding  17  full-length, truncated, or non-cleavable SRFs, and control LacZ. After 48 h, cell lysates were collected and analyzed for firefly and renilla luciferase activities using a dual luciferase reporter assay system (Promega) as per the manufacturer's instruction. The relative luciferase activities were normalized to the LacZ control group and presented as fold changes.  2.13 Affymetrix array analysis Microarray analysis of cardiac gene expression was performed as described previously (175). Hearts were harvested at 9 days following CVB3 infection and total RNA was isolated from mouse ventricular tissue using the RNeasy isolation kit (Qiagen) and pooled (n = 4 for each group). Samples were hybridized to a custom mouse GeneChip (Amgen) based on the Affymetrix GeneChip Expression Analysis Technical Manual (Affymetrix). In vitro transcription was performed using an ENZO Bioarray HighYield RNA Transcript Labeling kit as per the manufacturer's instructions. Arrays were washed in the Affymetrix Fluidics Station 450, stained with a streptavidin phycoerythrin solution and scanned using a HP Agilent GeneArray Scanner.  2.14 Real-time quantitative RT-PCR Total RNAs were extracted from murine HL-1 cardiomyocytes at 24 h post infection using RNeasy isolation kit. Reverse transcription was performed using SuperScript III Reverse Transcriptase (Life Technologies) according to the manufacturer’s instructions. mRNA levels of SRF and cardiac genes were measured by real-time quantitative PCR using an ABI Prism 7900HT Sequence Detection System as per the manufacturers' instruction (Perkin-Elmer Applied Biosystems) as previously described (34) and normalized to GAPDH mRNA.  2.15 Statistical analysis Two-way analysis of variance with multiple comparisons and paired Student's t-tests were performed. Data were shown as the mean plus minus standard deviation (S.D.). P-values < 0.05 were considered to be statistically significant. For statistical analysis of Affymetrix array data, the gene expression intensity values (averages of two replicates of the microarray analysis) were expressed as log2 ratios of CVB3- to sham-infected hearts and filtered for fold change cut-off of 1.8. Only genes that exhibited a 1.8-fold change (log2) or greater were included.  18  Chapter 3 – Autophagy in coxsackievirus infection 3.1  Background  3.1.1  Autophagy Autophagy, meaning self-eating, is a conserved process in eukaryotes by which cells removes  and recycles cytoplasmic long-lived proteins, misfolded proteins, as well as unwanted or damaged organelles to maintain cellular homeostasis under both normal and stress conditions (106, 109, 161). Autophagy is sub-classified in three forms: macroautophagy, microautophagy, and chaperone-mediated autophagy (42, 56, 67). Among them, macroautophagy, hereafter referred to as autophagy, is the most extensively studied form in both yeast and mammalian cells. 3.1.1.1 Process of autophagy The hallmark of autophagy is the formation of double-membrane vesicles called autophagosomes, which are derived from the elongation of crescent-shaped double-membrane structures known as isolation membranes or phagophores (106, 109, 161). The process of autophagy (depicted in Figure 3) takes place in four sequential steps: initiation, sequestration, fusion, and degradation. During the initiation step, the phagophore nucleates and elongates. Next, the phagophore sequestrates a portion of the cytoplasm including the target misfolded proteins or damaged organelles. The two ends of phagophore eventually fuse to form the autophagosome. Then, the outer membrane of the autophagosome fuses with the lysosomes to form the autolysosome. Finally, the inner membrane and the sequestrated contents of the autophagosome are degraded and recycled by hydrolytic enzymes. The recycled products (i.e. amino acids and fatty acids) are used for biosynthesis or other metabolic activities. Note that autophagosome may also fuse with early or late endosome to form amphisome prior to fusion with lysosome (60).  19  Lysosome Isolation membrane  Initiation  Autophagosome  Sequestration  Autolysosome  Fusion  Degradation  Figure 3. Process of autophagy pathway. The process of autophagy takes place in four sequential steps: initiation (formation of the isolation membrane), sequestration (elongation and closure of isolation membrane to sequestrate a portion of the cytoplasm), fusion (autophagosome-lysosome fusion), and degradation (formation of autolysosome to eliminate its inner membrane and sequestrated contents).  3.1.1.2 Autophagy pathway components The autophagy process is orchestrated by autophagy-related gene (ATG) products. More than 30 ATGs have been identified to date (83, 106, 146). The ATG proteins work together in complexes, including the class III phosphoinositide 3-kinase (PI3K) complex, the ULK complex, and two ubiquitination-like conjugation systems (ATG12-ATG5 system and LC3-PE (microtubule-associated protein light-chain 3-phosphatidylethanolamine) system) (83, 106, 146, 161). The ULK complex is made up of uncoordinated 51-like kinase 1/2 (ULK1/2), ATG13, focal adhesion kinase family interacting protein of 200 kD (FIP200), and ATG101. The class III PI3K complex is composed of vacuolar protein sorting-34 (Vps34), p150, beclin-1, ATG14, and Ambra1 (activating molecule in beclin-1-regulated autophagy). The two ubiquitination-like conjugation systems are comprised of ATG7, ATG12, ATG5, ATG16L1, ATG10, ATG4, ATG3, and LC3 (or ATG8 in yeast).  20  3.1.1.3 Molecular process of autophagosome formation The molecular process of autophagosome formation (Figure 4) is explained as follow. Autophagy is initiated with the activation of ULK complex, which in turn activates the class III PI3K complex (81, 97). Class III PI3K complex is translocated to the ER/trans-Golgi network (101, 211). It then helps the nucleation of isolation membrane by catalyzing the formation of phosphatidylinositol 3-phosphates (PI3Ps), which interact and recruit other ATGs involved in membrane elongation (184). The elongation of the isolation membrane is then mediated by the ubiquitination-like conjugation systems. In the ATG12ATG5 system, ATG12 is first activated by ATG7 and subsequently transferred to ATG10, then conjugated with ATG5 and subsequently to ATG16 to mediate the formation of LC3-PE (161). The LC3-PE system goes through a similar process in which nascent LC3 is first cleaved by ATG4 to become LC3-I, which is subsequently activated by ATG7, and then transferred to ATG3. Finally, LC3-I is conjugated to PE on the isolation membrane with the help of the ATG16-ATG12-ATG5 complex to form LC3-PE complex, also known as LC3-II (161). The recruitment of LC3-II on autophagic vesicles has been considered a common trait of autophagosome formation (161). Ratio of LC3-I to LC3-II, which measures the rate of LC3 lipidation, has also been widely accepted as a marker for autophagosome formation (98). 3.1.1.4 Regulation of autophagy The process of autophagy formation is tightly regulated by various kinases, phosphatases, and small GTPases. The serine/threonine protein kinase mTOR (mammalian target of rapamycin) is one key negative regulator of autophagy. Blocking mTOR activity by amino acid deprivation or rapamycin treatment has been shown to induce autophagy (41, 109). On the other hand, the eukaryotic initiation factor 2α (eIF2α) regulates autophagy positively (186). eIF2α is phosphorylated and activated in the event of starvation, endoplasmic reticulum (ER) stress, and the presence of double-stranded viral RNA replication-intermediates. eIF2α phosphorylation drives the formation of autophagosome as well as the inhibition of protein translation process (112, 186). In addition, small GTPases and phosphatases, such as Ras, PTEN, and protein phosphatase 2A, also participate in the regulation of autophagosome formation.  21  P  Growth factors ↓ Nutrients ↓ [ATP] ↓ [Ca2+] ↑ Rapamycin  Vps34 ULK  ATG101  Beclin-1  p150  ATG13 FIP200  2  1  Nucleation  ER Stress Starvation Double-stranded viral RNA  Beclin-1  mTOR  P  P  Vps34  eIF2α α  p150  Initiation  PE PI PI3P Other lipids  LC3/ ATG8 E2  ATG4  ATG3 LC3-I*  LC3-I ATP  LC3-II PE  ATP  ATG12  3  AMP + PPi  ATG7  ATG16 AMP + PPi  E1  * ATG12  Elongation  ATG12 ATG5  Atg10p  * = activated  E2 Isolation Membrane  Figure 4. Molecular machineries of autophagy pathway. Autophagy is initiated during cellular and metabolic stress via mTOR inactivation or eIF2α activation, which allows ULK complex activation. Activated ULK in turn phosphorylates Ambra1, which interacts with the Vps34 and Beclin-1 at the microtubule. Phosphorylation of Ambra1 results in the release of the class III PI3K complex from the microtubule and its translocation to the ER/trans-Golgi network, the major site of autophagosome formation. During autophagosome nucleation, the class III PI3K complex generates PI3Ps, which recruit other ATGs involved in membrane elongation. Elongation of the isolation  22  membrane is then mediated by two ubiquitination-like conjugation systems. ATG7 and ATG10 mediate ATG12-ATG5 complex formation, which is subsequently bound to ATG16. LC3 is cleaved by ATG4, forming LC3-I. Mediated by ATG7, ATG3, and the ATG16-ATG12-ATG5 complex, LC3-1 is conjugated to PE, generating LC3-PE complex, also known as LC3-II.  23  3.1.2  Autophagy in cardiac diseases Autophagy is vital for cardiomyocyte function and viability. Autophagy supplies emergency  energy sources by breaking down cytoplasmic proteins and organelles into amino acids and fatty acids. It was shown that cardiomyocyte survival under glucose deprivation is markedly reduced in the presence of autophagy inhibitor 3-methyladenine. Defects in the autophagy pathway lead to the development of cardiac diseases (121, 147, 190). It was reported that cardiac-specific ATG5 knockdown in adult mice results in cardiac hypertrophy, left ventricular dilatation, and contractile dysfunction (154). On the other hand, excessive autophagic activity induced under stress conditions such as ischemia reperfusion and pressure overloading is also detrimental to the heart as it leads into autophagic cell death (78, 134, 185, 197). This is supported by experimental data obtained from beclin-1 knockout mice challenged with either left anterior descending coronary artery ligation plus subsequent reperfusion or aortic banding (136, 216). In both models, disruption of beclin-1 expression results in the attenuation of stress-induced autophagy induction, as well as reduction in myocardial damage and pathological remodelling, underlining the damaging effect of dysregulated autophagy activity in the pathogenesis of various heart diseases. 3.1.3  Autophagy in viral infections Autophagy has also been suggested to play a part in host immune response by sequestrating  invading pathogens (e.g. herpes simplex virus and sindbis virus) and subsequently removing them from host cells (102). Viruses, on the other hand, have evolved various strategies to evade such host defense mechanism. Some viruses such as herpes simplex virus encode gene products that block cellular autophagy (186). Other viruses are able to hijack the autophagy pathway to benefit their own replication (102, 203). Certain positive-stranded RNA viruses such as poliovirus and coronavirus were reported to subvert cellular autophagy to facilitate their own replication (91, 171). Poliovirus proteins have been shown to co-localize with autophagic proteins (91). Induction of autophagy increases poliovirus yield, whereas autophagy inhibition decreases viral yield (91). Similar observations were made in murine hepatitis virus of the coronavirus family (171). Thus, the role of autophagy in the host seems to be dependent on the viral strain, but its overall relevance remains controversial.  24  3.2  Rationale Picornavirus infection induces reorganization of intracellular membrane, which serves as a  platform for the assembly of viral replication complex. However, the origin and nature of the viralinduced membrane platform remains unclear. Autophagosome, a double-membrane vesicle, is a potential site of viral replication. It has been reported that coronaviruses colocalize with the markers of autophagosome LC3 and ATG12 (171). Coronaviral replication is impaired in autophagy knockout (Atg5/-) embryonic stem cell lines (171). Additionally, poliovirus proteins have also been shown to colocalize with a co-transfected GFP-LC3 protein (91). Stimulation of autophagy increases poliovirus yield, and inhibition of the autophagosomal pathway decreases viral yield. However, the role of autophagy in CVB3 replication remains unknown. Here, I explored whether CVB3 manipulates the host autophagy protein degradation pathway to supply membrane platforms for successful viral replication.  3.3  Hypothesis and specific aims The objective of this chapter is to elucidate the molecular mechanisms utilized in viral  replication. In this study, my HYPOTHESIS is that coxsackievirus hijacks host’s cellular autophagy mechanism to facilitate its own replication. My SPECIFIC AIMS are: Aim 1. Examine whether CVB3 infection facilitates autophagosome formation Aim 2. Define the role of autophagosome in viral replication Aim 3. Identify the signalling components which stimulates the formation of autophagosome  25  3.4  Results  3.4.1  CVB3 infection increases autophagosome formation. To determine whether CVB3 infection triggers cellular autophagy, ultrastructural analysis was  performed with sham- or CVB3-infected HeLa and HEK293A cells. Electron micrographs of CVB3-infected cells revealed an increased prevalence of double-membrane vesicles in the perinuclear region compared to that of sham-infected cells (Figure 5A). Further quantitative analysis (Figure 5B) demonstrated a significant increase in autophagosome-like vesicles, which were characterized as double-membrane vesicles with a diameter of 0.2 to 1.0 μm present in the cytoplasm, in CVB3-infected cells. To confirm that the observed double-membrane vesicles were indeed related to autophagy, I examined LC3 modification, another hallmark of autophagy (107, 145). Two forms of LC3 have been reported. In quiescent cells, LC3 resides in the cytoplasm in a precursor form, referred to as LC3-I. However, upon exposure to various autophagy stimuli, it is rapidly activated and subsequently conjugated to phosphatidylethanolamine to form an LC3-phosphatidylethanolamine complex, commonly referred as LC3-II, which participates in the formation of autophagosomes and remains attached to matured autophagosomes (98). It has been shown that the LC3-II/LC3-I ratio correlates well with the number of autophagosomes (98). After CVB3 infection, Western blots were performed to detect the LC3-II/LC3-I ratio, using an anti-LC3 antibody which recognizes both forms of LC3. As shown in Figure 5C, expression of LC3-II increased with the progression of CVB3 infection, which was accompanied by a decrease of LC3-I expression. The densitometry ratio of LC3-II to LC3-I demonstrated a cumulative increase in autophagosomes during CVB3 infection. Punctate accumulation of LC3, which represents the recruitment of LC3 to autophagic vacuoles, has also been suggested to be a characteristic of autophagy (98). After CVB3 infection, I showed that GFP-LC3-transfected cells with punctate GFP-LC3 distribution were markedly increased (Figure 5D), further confirming the increased formation of autophagosomes after viral infection.  26  27  Figure 5. CVB3 infection increases autophagosome formation. (A) Representative electron micrographs of sham- and CVB3-infected HeLa (top) and HEK293A (bottom) cells at 7 h post infection. Arrows indicate representative autophagosomes that would be scored positive in panel B. (B) Quantitation of the numbers of autophagosomes in sham- and CVB3-infected HEK293A cells. Data shown represent the number of autophagosomes per cell under each condition (mean ± standard error [SE]). #, P < 0.001 compared to sham infection, by Student's t test. (C) The LC3II/LC3-I ratio, which is a hallmark of autophagy, increases with the progression of CVB3 infection. (Left) Western blot analysis of LC3 and viral capsid protein VP1 expression in CVB3-infected cells at various times post infection. β-Actin expression was examined as a protein loading control. (Right) Expression of LC3-II and LC3-I and ratio of LC3-II to LC3-I at the indicated times after CVB3 infection (means ± SE; n = 5 or 7 [as indicated]). LC3 expression was quantitated by densitometric analysis using NIH ImageJ v1.37  28  and normalized to sham infection, which was arbitrarily set to a value of 1.0. and, P < 0.01 compared to sham infection. (D) Representative confocal images and quantitation of autophagosomes in GFP-LC3transfected HEK293A cells after CVB3 infection. HEK293A cells were transfected with GFP-LC3 plasmid for 24 h, followed by CVB3 infection. The GFP fluorescence was analyzed by confocal fluorescence microscopy. The percentage of cells with punctate GFP-LC3 localization relative to all GFP-positive cells was calculated and is presented as the mean ± SE (n = 5). #, P < 0.001 compared to sham infection.  29  3.4.2  Mechanism of induction of autophagosome formation To determine the potential mechanism of induction of autophagosome formation, I examined  the phosphorylation status of eIF2α and mTOR, two key regulators that control autophagosome formation and maturation. It has been shown that eIF2α phosphorylation positively regulates the autophagic pathway (186), whereas mTOR is a negative regulator of autophagosome formation (41). I found that eIF2α phosphorylation was increased at 5 and 7 h post infection (Figure 6), corresponding to an increased formation of autophagosomes in CVB3-infected cells. However, no significant changes in the levels of mTOR phosphorylation were observed after CVB3 infection (data not shown).  Figure 6. CVB3 infection induces eIF2α phosphorylation. Western blot analysis of phosphorylated eIF2α and β-actin (loading control) expression was performed at the indicated times post infection. Similar results were obtained in two independent experiments.  30  3.4.3  Autophagy inhibitor 3-MA reduces CVB3 replication. To determine whether the autophagosome induction during CVB3 infection was a host antiviral  response or a viral replication mechanism, I tested the effect of 3-methyl adenine (3-MA) on CVB3 replication. 3-MA is a widely used selective inhibitor of autophagy which blocks the formation of autophagosomes and inhibits intracellular protein degradation without affecting protein synthesis (178). As shown in Figure 7, treatment with 3-MA resulted in a dose-dependent reduction of viral capsid protein (VP1) expression (Figure 7A) as well as in a significant reduction in CVB3 viral progeny titer (Figure 7B). More importantly, I found that 3-MA treatment led to significant inhibition of viral protein synthesis, even when it was introduced during the later stages of the virus life cycle (Figure 7C). This finding suggests that autophagic machinery may not have a destructive role during CVB3 infection and instead may play an important role in the replication of CVB3.  31  Figure 7. Autophagy inhibitor 3-MA reduces CVB3 replication. (A) (Top) Western blot analysis of viral protein VP1 and β-actin (loading control) expression 7 h following CVB3 infection, with or without 3-MA treatment, as indicated. (Bottom) VP1 expression was quantitated by densitometric analysis using NIH ImageJ v1.37 and normalized to β-actin expression. The VP1/β-actin ratio was further normalized to the CVB3-infected group without treatment, which was arbitrarily set to a value of 1.0. The data shown are means ± SE (n = 3), and significance was determined by Student's t test. *, P < 0.05; #, P < 0.001 (compared to nontreated cells). (B) Plaque assay measuring the CVB3 viral progeny titers in supernatants collected from infected cells treated with or without 3-MA. The data shown are means ± SE for three independent experiments. *, P < 0.05 compared to nontreated cells. (C) Western blot analysis of CVB3-infected cells treated with 10 mM 3-MA at different times, as indicated.  32  3.4.4  Induction of autophagy enhances viral growth. To further determine the role of autophagy, I investigated the effect of autophagosome  induction on viral protein expression. Cells were deprived of amino acids (Figure 8A), a condition that is known to be able to induce autophagy (41), or treated with the pharmacological reagent rapamycin, which has been shown to induce autophagy by inhibiting the mTOR pathway (23) (Figure 8B). I showed that induction of autophagy through either nutrient deprivation or rapamycin treatment significantly increased viral protein expression (Figure 8). These results further suggest a critical role of autophagy in the virus life cycle, although I cannot rule out other effects that these treatments might have on viral replication, independent of autophagy.  33  Figure 8. Induction of autophagy enhances viral replication. Western blot analysis of VP1 and β-actin expression was performed at 7 h post infection. (A) Prior to CVB3 infection, cells were incubated in Hank's balanced salt solution for 2 h to induce cell starvation. (B) Cells were pretreated with rapamycin (50 nM) for 3 h. VP1 expression was quantitated, normalized, and analyzed as described in the legend to Figure 7 (data are means ± SE; n = 3). *, P < 0.05; #, P < 0.001 (compared to nontreated cells).  34  3.4.5  Knockdown of the genes critical for autophagosome formation reduces CVB  replication. To extend the studies with pharmacological agents, I employed a target-specific RNA interference approach to assess the effect of autophagosome disruption on viral replication. Cells were transfected with siRNAs (for ATG7, Beclin-1, and Vps34) to knock down the genes required for different steps of autophagosome formation. The autophagy protein ATG7 is an enzyme essential for activation of autophagosome formation and maturation machinery (106, 109). Deletion of ATG7 in mice has been shown to result in reduced autophagosome formation and impaired degradation of proteins and organelles induced by starvation (111). As shown in the phase-contrast images in Figure 9A, cells transfected with siRNA against ATG7 displayed greater resistance to CVB3 infection and maintained normal cell morphology, unlike the scramble siRNA-treated cells, which displayed severe virus-induced cytopathic effect. In addition, ATG7 siRNA treatment also blocked viral protein synthesis (Figure 9B). Similarly, disruption of the class III PI3K signaling complex required for autophagosome formation by Beclin-1 and Vps34 siRNAs resulted in significant reductions in CVB3 replication (Figure 10). My results further demonstrate a requirement of autophagy for effective coxsackievirus infection.  35  Figure 9. Gene silencing of ATG7 by siRNA inhibits CVB3 replication. (A) Representative light microscopic images of CVB3-infected cells, with or without ATG7 siRNA transfection. (B) Western blot probing for VP1 and ATG7 expression, with or without ATG7 knockdown. VP1 expression was quantitated, normalized, and analyzed as described in the legend to Figure 7 (data are means ± SE; n = 3). *, P < 0.05 compared to scramble siRNA-transfected cells.  36  Figure 10. Knockdown of class III PI3K components reduces CVB3 replication. (A) Western blot analysis of VP1 and Beclin-1 or Vps34 expression in CVB3-infected cells transfected with Beclin-1 (left) or Vps34 (right) siRNA prior to CVB3 infection. VP1 expression was quantitated, normalized, and analyzed as described in the legend to Figure 7 (data are means ± SE; n = 3). *, P < 0.05 compared to scramble siRNA control. (B) Plaque assay results for scramble or Beclin-1 and Vps34 siRNAtransfected cells (means ± SE; n = 3). *, P < 0.05; #, P < 0.001 (compared to scramble siRNA-transfected cells). 37  3.4.6  Expression of p62 following CVB3 infection. I next examined the effect of CVB3 infection on autophagy- or lysosome-mediated protein  degradation. The protein p62 has been considered a marker for autophagy-mediated protein degradation or autophagic flux (108). As shown in Figure 11, inhibition of lysosome function by bafilomycin A and NH4Cl, two pH-neutralizing agents, resulted in increased expression of p62. However, protein expression of p62 was unchanged during the course of CVB3 infection, suggesting that CVB3 infection triggers increased accumulation of autophagosomes without promoting protein degradation by lysosomes.  A -  NH4Cl BFLA p62 β -actin  B CVB3 sham  1h  3h  5h  7h p62 VP1 β -actin  Figure 11. Expression of p62 is not affected by CVB3 infection. (A) Western blot analysis of p62, a marker of autophagic protein degradation activity, in cells treated with the lysosome inhibitors bafilomycin A (BFLA; 0.1 μM) and NH4Cl (2.5 mM) for 16 h. (B) Expression of p62 in CVB3-infected cells at the indicated times after virus infection. Similar results were obtained in two independent experiments.  38  3.4.7  Lysosomal inhibition enhances CVB3 replication. Finally, I investigated whether prevention of autophagosome-lysosome fusion promotes viral  infectivity. Cells were transiently transfected with scramble or LAMP2 (lysosome-associated membrane protein 2) siRNA for 48 h and then infected with CVB3. I showed that knockdown of LAMP2 resulted in significant increases of viral protein expression and virus titer (Figure 12). LAMP2 is a lysosomal membrane protein critical for autophagosome-lysosome fusion. Knockdown of LAMP2 has been shown to induce the cytoplasmic accumulation of LC3-associated vesicles (75). It was also reported that LAMP2-deficient mice exhibit a massive accumulation of autophagic vacuoles in various tissues (188). Thus, my results further suggest a key role of autophagosomes in supporting CVB3 replication.  39  Figure 12. Lysosomal inhibition enhances CVB3 replication. (A) Western blot analysis of VP1 and LAMP2 expression in CVB3-infected cells transiently transfected with scramble or LAMP2 siRNA. VP1 expression was quantitated, normalized, and analyzed as described in the legend to Figure 7 (data are means ± SE; n = 3 or 4 [as indicated]). (B) Virus titer in scramble or LAMP2 siRNA-transfected cells, determined by plaque assay (mean ± SE; n = 3). *, P < 0.05 compared to scramble siRNA control.  40  3.5  Discussion Autophagy is an evolutionarily conserved cellular process that spans the entire spectrum of  eukaryotes. It serves a housekeeping role in protein turnover and in the removal of damaged or unnecessary organelles. It also serves an important function in innate host defense by eliminating intracellular pathogens. For example, it has been shown that pharmacological induction of autophagy helps in the clearance of Mycobacterium tuberculosis (79) and that CD40 induction of autophagy promotes elimination of Toxoplasma gondii from macrophages (7). Furthermore, recent studies suggest a role for autophagy in host adaptive immune responses through the delivery of intracellular pathogenic antigens for major histocompatibility complex class II presentation to CD4 T cells (176). Successful invading pathogens have evolved various strategies to evade host autophagy. For instance, some bacterial or viral proteins, such as IcsB secreted by Shigella flexneri (160) and ICP34.5 encoded by herpes simplex virus type 1 (186), have been documented to block or evade cellular autophagy. Other pathogens adopt different approaches in battling host autophagic defense by subverting it to facilitate their own procreation. In fact, several studies have shown that the autophagosome provides an intracellular compartment that benefits the survival and multiplication of bacteria, including Porphyromonas gingivalis, Brucella abortus, and Coxiella burnetii (176). In addition, some viruses have been reported to induce autophagosome formation to facilitate their own replication (102, 203). Replication of positive-stranded RNA viruses requires intracellular membrane surfaces on which they assemble their replication complexes. Colocalization of viral RNA replication machineries with double-membrane vesicles in infected cells has been well documented for various positive-stranded RNA viruses, including poliovirus, rhinovirus, equine arterivirus, murine hepatitis virus, and severe acute respiratory syndrome virus (59, 203). Thus, it was speculated that the autophagosome, a doublemembrane vesicle, may serve as a site for viral RNA replication. Recent studies on coronavirus and poliovirus seem to support this notion. It was reported that murine hepatitis virus, which belongs to the coronavirus family, colocalizes with the ubiquitin-like autophagy proteins (LC3 and Atg12p) and that viral replication is impaired in autophagy knockout (Atg5−/−) embryonic stem cell lines (171). Additionally, poliovirus proteins have been shown to colocalize with a co-transfected GFP-LC3 protein (91). Stimulation of autophagy increases the poliovirus yield, and inhibition of the autophagosomal pathway decreases the viral yield (91). Despite this exciting observation, recent data on the role of autophagy in positive-stranded RNA virus infection remain contradictory. A recent report showed that infection of rhinovirus type 2, a member of the picornavirus family, does not induce autophagosome formation (25).  41  That paper further reported that neither induction nor inhibition of autophagy affects viral protein production. In addition, using ATG5-deficient cells, it was recently demonstrated that ATG5 and an intact autophagic pathway are not required for the replication of murine hepatitis virus (214). These studies challenge the view that the autophagosome is a common cellular component utilized by positivestranded RNA viruses to facilitate their replication. These discrepancies of the reported data prompted me to explore the role of autophagy in coxsackievirus infection in this study. I report here that CVB3 infection induces the formation of doublemembrane vesicles with concurrent LC3 lipidation. I further showed that manipulation of the host autophagy pathway by pharmacological compounds or target-specific RNA interference resulted in significant alterations in the outcome of CVB3 replication. I demonstrated that inhibition of signaling pathways or autophagy genes critical for autophagosome formation reduces viral protein expression and the viral progeny titer. Conversely, increasing the cellular number of autophagosomes prior to viral infection by cellular starvation, rapamycin treatment, or lysosomal inhibition promotes viral replication. Taken together, my data suggest that the host double-membrane autophagosome is likely utilized by CVB3 to facilitate its own replication. Of note, autophagy activation is most prominent at the later stages of viral replication. Increased formation of autophagosome helps the exponential amplification of viral components by supplying sufficient physical scaffolds for the viral components synthesized at earlier time points. As alluded to earlier, the process of autophagosome formation is tightly controlled. The serine/threonine protein kinase mTOR is one of the key regulators and negatively controls autophagosome formation (41). Class I PI3K is the traditional upstream activator of mTOR. In addition, it has been shown that mTOR can be regulated by intracellular amino acids and ATP, independent of class I PI3K activation. ATP depletion or specific inhibition of mTOR by rapamycin has been shown to activate autophagy, whereas the addition of amino acids inhibits this process. In this study, I showed that levels of mTOR phosphorylation were unchanged after CVB3 infection, suggesting that CVB3-induced autophagy is not triggered by mTOR dephosphorylation. In contrast, eIF2α kinase positively regulates autophagy in response to nutrient deprivation and viral infection. eIF2α kinase plays a key role in translation inhibition under various stress conditions, through phosphorylation of eIF2α. Restriction of herpes simplex virus growth has been linked to such a function of eIF2α phosphorylation (149). Recently, the eIF2α kinase signaling pathway was also shown to promote autophagy induced by nutrient deprivation or rapamycin treatment, likely through a mechanism that induces LC3 conversion (112, 186). My present study demonstrates that eIF2α is  42  phosphorylated after CVB3 infection, suggesting a potential signaling mechanism for CVB3-induced autophagosome formation. It is unclear how eIF2α phosphorylation occurs during CVB3 infection. One explanation could be that double-stranded RNA formed from a CVB3 intermediate activates doublestranded RNA-dependent protein kinase and subsequently leads to eIF2α phosphorylation. Class III PI3K complexes also play a crucial role in the early steps of autophagosome formation. In mammalian cells, the autophagic protein Beclin-1 (a homologue of yeast ATG6) forms the class III PI3K complex with Vps34, a class III PI3K, at the trans-Golgi network (101, 211). This complex promotes autophagosome formation by supplying phosphatidylinositol 3-phosphates to the preautophagosome membrane (184). It has been shown that 3-MA blocks the formation of autophagosomes by inhibiting the activity of class III PI3K (23, 169). I demonstrate that 3-MA blocks CVB3 infection, which is in agreement with previous report from my laboratory that inhibition of PI3K by LY294002 suppresses CVB3 replication (57). LY294002 is an inhibitor of both class I and class III PI3K and has also been shown to be a potent autophagy inhibitor (23). Although the specific mechanism by which the autophagosome is utilized during coxsackievirus replication in host cells has not been elucidated, it is likely that, as demonstrated for poliovirus (91), the autophagosome provides a physical scaffold where the coxsackieviral complex may reside and viral RNA synthesis may occur. Furthermore, LC3 has been identified to be a binding protein of an AU-rich RNA element in the 3’-untranslated region of fibronectin mRNA, which facilitates the recruitment of RNA to polyribosomes for translation (215). Perhaps AU-rich element-containing viruses, such as CVB3, which carries an AU-rich element in the 3’-untranslated region (170), also adopt LC3-docking doublemembrane vesicles as the sites of viral protein translation through the recruitment of polyribosomes. It is plausible to speculate that autophagosome double-membrane vesicles not only act as sites of viral RNA synthesis but also take an active role in CVB3 protein synthesis by bridging the nascent viral RNA with polyribosomes through LC3.  3.6  Limitations and solutions Multiple experimental methods were conducted in my study to ensure the validity of each  finding. For example, I confirmed the induction of autophagosome formation in CVB3 infected cells by transmission electron microscopy and quantification of LC3 conversion. I increased the level of autophagosome in host cells by nutrient starvation, rapamycin treatment, and lysosomal inhibition. I inhibited the autophagy pathway using inhibitor (3-MA) against the Class III PI3K, as well as siRNA against autophagy component ATG7. However, several supplemental experiments could further  43  strengthen my observations. I could apply immune-gold staining of LC3 to demonstrate the presence of LC3 on CVB3-induced vesicles observed in transmission electron microscopy. I could also monitor the autophagic flux, i.e. the turnover of autophagosome, in CVB3 infected cells. By comparing CVB3-infected cells with or without the introduction of lysosomal inhibitors (e.g. bafilomycin A, NH4Cl, and choloroquine) an hour prior to cell harvesting, I can gain further insights in the nature of virus-induced autophagosomes. The difference in LC3 conversion between these treatment groups demonstrates whether the autophagosomes mature into autolysosomes or they stay as is. So far, my p62 expression result suggests that the virus-induced autophagosomes are not geared towards degradation. Measuring the autophagic flux would provide a more definitive answer. Regarding the autophagy inhibition experiments, additional experiments could help validate my observations. In this study, I only achieved successfully knockdown ATG7 expression using siRNA. I tried other siRNAs against ATG5 and LC3, but failed to observe any meaningful knockdown in western blot confirmation. Higher efficiency knockdown of other autophagy components using lentivirus shRNAs or ATG knockout cell lines may help elucidate the specific requirements of autophagy pathway in CVB3 infection. It is possible that CVB3 only utilize part of the autophagy machineries, therefore ATG knockdown beyond the ubiquitination-like conjugation systems (ATG12-ATG5 and LC3-PE) such as the ULK complex components may provide further clarifications. Autophagy inhibitor 3-MA is particular potent in suppressing CVB3 replication (Figure 7C). It effectively inhibited CVB3 protein expression even when introduced at 5 hours post infection. However, 3-MA may have other properties in addition to autophagy inhibition. More specific inhibitors targeting the ubiquitination-like conjugation system should also be used. I did however perform siRNA knockout of the components (Vps34 and beclin-1) of Class III PI3K, the target of 3-MA. Knockdown of these genes also inhibited CVB3 replication. Knockdown of autophagy components linking Class III PI3K to the rest of the autophagic machineries will help further elucidate the role of this complex in CVB3-infection. Another limitation of the present study is the utilization of tumor cell lines, HEK293 (transformed Human embryonic kidney cells) and HeLa (Human cervical cancer cells). These cell lines are great for the study of molecular pathways due to their ease of cultivation and transfection. However, cardiomyocytes such as rat neonatal cardiomyocytes and HL-1 cells should be used to confirm the findings in this chapter.  44  3.7  Future directions In this study, I demonstrated that eIF2α, which is a known positive regulator of the autophagy  pathway, is phosphorylated during CVB3 infection. However, it is by no means a conclusive mechanism of autophagy subversion. Future studies are required to clarify the relationship between eIF2α phosphorylation and autophagy induction in CVB3 infection. In addition, the mechanism behind eIF2α phosphorylation in CVB3 infection remains to be explored. Perturbation of its upstream kinases such as dsRNA-dependent protein kinase R (PKR) and protein kinase RNA-like endoplasmic reticulum kinase (PERK), as well as phosphatases such as protein phosphatase 2A and PTEN using chemical inhibitors or RNA interference may reveal the molecular signalling events that lead to autophagosome formation in CVB3-infected cells. Coxsackieviral proteins are potential molecular drivers of autophagosome formation. As previously described, non-structural proteins 2B, 2BC, and 3A are involved in intracellular membrane rearrangement. 3A plays an important role in membrane trafficking and in manipulating intracellular protein transport from ER to Golgi (200). Mutations in 3A result in defects in viral RNA synthesis (200, 201). Physical association of 3A and viral polymerase 3D has also been shown to play an important role in the initiation of viral RNA synthesis (85). 2B and its precursor 2BC alter membrane permeability (3, 35, 194). 2B was also reported to have anti-apoptotic properties that prevent premature cell death, so as to ensure sufficient time for viral replication (193). However, the exact mechanisms by which 3A, 2B, and 2BC regulate intracellular membrane reorganization are still largely unclear. Future studies correlating the expression and localization of these CVB3 non-structural proteins (alone or in combinations) with the markers of autophagosome, as well as identifying their cellular binding partners may help elucidate the role of these membrane-modifying viral proteins in autophagy subversion. As described above, LC3 has been shown to bind the AU-rich elements (AUUUA sequence) in the 3’UTR of fibronectin mRNA and facilitate the recruitment of polyribosomes for translation (215). CVB3 genome also contains AU-rich elements in its 3’UTR (170). CVB3’s 3’UTR has been shown to play a critical role in viral replication by modulating mRNA stability and translation efficiency (103, 209). I therefore postulated that the autophagosome, harbouring LC3, is an ideal platform for CVB3 replication because it may support the concurrent translation of newly synthesized viral RNA through the recruitment of polyribosomes. Future studies involving the co-localization of viral RNA-dependent RNA polymerase 3D with markers of autophagosome and polyribosome, as well as mutation of AU-rich elements in the 3’UTR of CVB3 genome may help test the speculation.  45  My in vitro observation of autophagy subversion by coxsackievirus warrants further investigation in vivo. Confirmation of autophagosome formation in vivo is made feasible by the establishment of GFP-LC3 transgenic mice (148). CVB3 infection of GFP-LC3 mice will elucidate the dynamics of autophagy activation in the progression of viral myocarditis. In fact, preliminary observations from my coworkers (unpublished data) and others (99) have demonstrated the activation of autophagy in CVB3-infected heart, liver, and pancreas using GFP-LC3 mice. Further studies using cardiac-specific autophagy knockout mice will reveal the role of autophagy in the pathogenesis of viral myocarditis. My study demonstrated that gene-silencing of ATG7 in cells by siRNA inhibits CVB3 infection. ATG7 is an enzyme essential for both ubiquitination-like conjugation systems required for autophagosome formation and maturation (106, 109). Knockdown of ATG7 was reported to be more efficient in blocking autophagy pathway than the knockdown of other ATGs. Deletion of ATG7 in mice was reported to result in attenuation of autophagosome formation and impaired degradation of proteins and organelles induced by starvation (111). Therefore, cardiac-specific disruption of autophagy pathway through ATG7 knockout in mice will be ideal in studying the role of autophagy in the overall development of viral myocarditis.  3.8  Summary I demonstrated that CVB3 manipulates the host autophagy protein degradation pathway to  supplement viral replication. Autophagy is an evolutionary conserved homeostatic mechanism in eukaryotes that degrades and recycles long-lived cytoplasmic proteins, as well as damaged organelles. The hallmark of autophagy is the formation of double-membrane vesicles known as autophagosomes. I provided the initial evidence that CVB3 infection induces the formation of autophagosomes. Upregulation of autophagosome formation enhances CVB3 replication, whereas downregulation of autophagy pathway reduces CVB3 replication in host cells. My results help clarify the nature of the intracellular membranes previously shown to be required for viral replication.  46  Chapter 4 – Serum response factor in CVB3-induced myocarditis 4.1  Background  4.1.1  Mechanisms of CVB3-induced DCM Although viral myocarditis was originally considered an immune system-mediated disease of the  heart, accumulating evidence demonstrates that early direct virus-mediated injury occurring prior to infiltration of immune cells contributes significantly to the pathogenesis of viral myocarditis (37, 86, 138). In cultured cells, CVB3 infection induces a direct cytopathic effect and cell apoptosis (29, 30, 33). CVB3 infection of severe combined immune deficient mice results in early and extensive myocyte coagulative necrosis and contraction band necrosis as compared to their wild-type counterparts (37, 138). Prominent cytopathic alterations co-localize to cells which desmonstrate viral replication by in situ hybridization for viral RNA, reinforcing the importance of direct virus-induced damage (105). Viral proteases have been recognized as important pathogenic factors contributing to the development of DCM. It has been shown that transgenic mice with inducible, cardiac-specific expression of viral protease 2A develop severe cardiac dysfunction and DCM (205). Experimental evidence from cell culture and cell-free in vitro studies demonstrates that 2A induces the cleavage and functional impairment of dystrophin (11-13). As depicted in Figure 13, dystrophin connects the cytoskeletal actinbinding site to the β-dystroglycan extracellular matrix anchor; thus, its cleavage leads to the disruption of the cytoskeletal architecture. It is therefore proposed that 2A-induced DCM is associated with disrupted myocyte membrane integrity through the cleavage of dystrophin (11-13). However, a direct causal relationship between dystrophin cleavage and 2A-induced DCM has not been provided in these studies and other mechanisms may also play pivotal roles. Moreover, mdx (dystrophin-deficient) mice display a relatively mild DCM phenotype (50, 76). The mild phenotype has been attributed to the compensatory up-regulation of the dystrophin homologue, utrophin, as evidenced by severe dystrophic phenotypes in mice with double-knockout of utrophin and dystrophin (50, 76, 80, 93). Previous studies have shown that CVB3 infection does not cleave utrophin (13). Thus, the DCM phenotype induced by dystrophin cleavage may be dampened by the compensatory effect of utrophin. Together, these data suggest that dystrophin-cleavage alone is not sufficient to explain the severe cardiomyopathy phenotype observed in 2A transgenic mice. Thus, I postulate that other substrates of 2A may also play a key role in the development of DCM.  47  Sarcospan Sarcoglycans  α-dystroglycan β-dystroglycan  Sarcolemma  2A Syntrophins Cleaved dystrophin  Dystrophin  Actin cytoskeleton  Increased sarcolemmal permeability  Decreased force transmission  Myocyte dysfunction  Figure 13. Coxsackieviral protease 2A cleaves dystrophin at its 3' hinge. Dystrophin is a component of the dystrophin-glycoprotein complex that links the cytoskeleton to the extracellular matrix. Dystrophin cleavage by viral protease 2A contributes to myocyte dysfunction by reducing contractile force transmission and increasing sarcolemmal permeability.  48  4.1.2  Serum response factor Serum response factor (SRF), originally identified by Treisman and colleagues, is a founding  member of the MADS (MCM1 in yeast, Agamous and Deficiens in plants, and SRF in animals) box family of transcription factors (159, 191). SRF was named after its binding activity to serum response element (SRE, also known as CArG box (CC(A/T)6GG)) located in the promoter region of immediate early transcription factor, c-Fos (159, 191). Similar SREs were subsequently found in the promoter regions of a number of other immediate early genes, as well as contractile genes and microRNA (miRNA) genes (143, 157, 162). A genome-wide search for SRF targets has identified approximately 1 200 genes, which contain single or more copies of this consensus SRE sequence in their promoter regions (termed CArGome) (183). More than 250 of these genes have been validated (183). 3.1.2.1 Important SRF domains Gene expression regulation property of SRF is conferred by its major domains (see Figure 14) (31, 143, 162). Its nuclear localization signal (amino acids 95-100) is required for nuclear translocation of SRF. Its DNA binding and dimerization domain (amino acids 133-222) is highly conserved among eukaryotes and is essential for its binding to SREs of target genes, as well as formation of SRF dimers. Its transactivation domain (amino acids 339-508) carries out transcriptional activation of target genes. It also has multiple phosphorylation sites targeted by several signaling kinases. 3.1.2.2 SRF regulation SRF by itself has relatively low transcriptional activity. Expression of SRF target genes are controlled in a cell- and context-specific manner through interactions between SRF and its cell-specific cofactors. Examples of SRF cofactors in cardiac tissues are Nkx2.5, GATA4, MEF2, and Myocardin. SRF cofactors may compete for common binding interface on SRF, thus allowing SRF-dependent gene expression in varying contexts. SRF-mediated gene expression can be activated by a variety of agents, including serum, cytokines, mitogens, lipopolysaccharides, and viral activator proteins. All in all, the activity of SRF is governed by various mechanisms including transcription, translation, alternative RNA splicing, association with cofactors, subcellular localization, as well as phosphorylation and cleavage (31, 143, 162).  49  DNA binding/dimerization NLS domain (MADS box) SRF  Transcriptional activation domain  N  C 1  95 100 133  222  339  508 aa  Cardiac regulatory proteins GATA4, Nkx2.5, ANF…  SRF  SRF  Sarcomere contractile proteins β -MHC, MLC, α -actin… Cardiac-enriched miRNAs miR-1, miR-2, miR-133, miR-486… SRF target genes in heart Transcription  SRE  SRF Target Genes  SRE = CArGSRF box Co-factors SRE = CArG box  Figure 14. Serum response factor regulates cardiac gene expression. SRF is a muscle-enriched transcription factor that regulates the expression of cardiac regulatory proteins, sarcomere contractile proteins, as well as cardiac-specific microRNAs (miRNAs). SRF contains three major domains: NLS (nuclear localization signal), DNA binding and dimerization domain, and transcriptional activation domain. SRF dimerizes and associates with co-factors such as GATA4, Nkx2.5, MEF2, and myocardin and binds to serum response element (SRE) (also known as CArG box) to activate gene transcription.  50  3.1.2.3 SRF in cell growth and differentiation SRF is an important regulator of cell growth, differentiation, as well as apoptosis. Blocking SRF function by anti-SRF antibodies or by SRF sequence-specific antisense RNA decreases muscle marker gene expression and prevents the differentiation of myoblast to myotube (73, 182). In contrast, activation of SRF through the Rho-GTPases pathway promotes muscle cell differentiation and muscle gene expression (28). It has been demonstrated that caspase-3 activation leads to the cleavage of SRF in human B-cells, whereas the expression of a caspase-resistant form of SRF significantly inhibits apoptosis in these cells (21, 53). Knockout of SRF gene in mice is embryonic lethal due to defects in mesoderm differentiation (10). 4.1.3  SRF-mediated cardiac gene expression SRF is highly expressed in cardiac muscle and plays a central role in cardiac development and  function by regulating genes for cardiac contractile and regulatory proteins as well as silencer miRNAs (124, 144, 155, 156). Cardiac muscle-specific inactivation of SRF in utero results in a dilated, heart failure-like phenotype with embryo demise occurring at ~e11.5 of mouse development (144, 156, 158, 166). Cardiac genes directly under SRF regulation include cytoplasmic cytoskeletons such as desmin and titin, membrane cytoskeletons such as dystrophin and zona occludens, contractile elements such as cardiac α-actin, α-myosin heavy chain (α-MHC), β-MHC, myosin light chain (MLC), and cardiac troponins, contractile regulators such as sarcoplasmic reticulum Ca2+-ATPase (SERCA), Na+/Ca2+ exchanger NCX1, and atrial naturetic factor (ANF), as well as transcription regulators such as nuclear factor of activated Tcells (NFAT), Nkx2.5, myocardin (144, 156, 158, 166, 183). Indirect SRF regulation of cardiac genes has also been reported. Transcription factors with no functional SRE in their promoter regions, such as GATA-4 and myocyte enhancer factor-2 (MEF2), are also reduced in SRF-null hearts and cardiomyocytes (144, 156, 158, 166). Downregulation of these indirect targets is suggested to be secondary to the reduced expression of other SRF targets. Interestingly, SRF-inactivation in cardiomyocytes also activates a subset of genes, which is attributed to the loss of SRF-dependent miRNAs, illustrating an additional gene-silencing function of SRF (14, 156). Recent studies have identified several miRNAs as targets of SRF in the heart, which include miR-486 (180), miR-133a (124, 156), and miR-1 (213). Expression of these miRNAs controls heart development and function by repressing the expression of cardiac regulatory proteins.  51  4.1.4  SRF in heart diseases Severe heart failure is associated with the reduced expression of a broad range of cardiac-  specific genes. Microarray studies in human end-stage heart failure revealed a significant downregulation of many cardiac genes, including many SRE-containing genes, implicating the importance of SRF in the progression of heart failure (17, 90, 187). It was reported that expression of a dominant-negative splice variant of SRF that lacked portions of the transactivation domain is markedly elevated in human and animal failing hearts (20, 46). The dominant-negative SRF splice variant potently inhibited SRF-dependent gene expression, mirroring the biochemical phenotype seen in SRF-null mice. Interestingly, a recent report showed that SRF cleavage also occurs in the cardiomyocytes of end-stage DCM patients (32). Increased caspase activity during heart failure induces the cleavage of SRF, producing a truncated protein that lacks the C-terminal transactivation domain, and acts as a dominant repressor of SRF-dependent cardiac genes, similar to the alternatively spliced variant (32). Together, these studies suggest that decreased expression of full-length wild-type SRF and/or the increased production of truncated SRF may down-regulate SRF-dependent genes and thereby contribute to the progression of severe heart failure. On the other hand, sustained elevation of SRF expression is also detrimental to the heart. SRF overexpression induces cardiac hypertrophy, which is a compensatory response to a variety of physiological or pathological stimuli. Re-expression of fetal genes such as ANF, β-myosin heavy chain, and skeletal α-actin, has been considered as a marker of this condition (2, 15, 153). Many of these genes have SREs in their promoter regions suggesting that SRF may be a critical transcription factor in the reprogramming of fetal genes. In vitro experiments using isolated cardiomyocytes from SRF-floxed transgenic mice demonstrated the requirement of SRF in the induction of fetal gene expression (155). Transgenic mouse studies showed that cardiac-specific overexpression of SRF induces a pattern of fetal gene re-expression similar to that observed in cardiac hypertrophy, and subsequently causes severe cardiomyopathy characterized by four-chamber dilatation, cardiomyocyte hypertrophy, interstitial fibrosis, and mitochondrial/myofibril damage (212). Most recently, there was a report showing that forced expression of striated muscle activator of Rho signalling sensitizes the heart to pressure overloadinduced and calcineurin signalling-induced cardiac hypertrophy by modulating SRF-dependent transcription (113). Collectively, these experimental data suggest that strictly-controlled SRF expression is essential for the maintenance of cardiac homeostasis.  52  4.2  Rationale As described previously, enteroviral-encoded proteases have been recognized as important  pathogenic factors contributing to the development of DCM. Transgenic mice with cardiac-restricted expression of viral protease 2A develop severe cardiac dysfunction and DCM (205). Disruption of the dystrophin-glycoprotein complex through dystrophin cleavage by 2A has been proposed as one important mechanism for the progression of DCM (11-13). However, dystrophin-null mice do not develop severe cardiomyopathy (50, 76) as observed in 2A transgenic mice, suggesting that other molecular targets for the protease exist and contribute to the viral pathogenesis. In addition, in-house Affymetrix array data (Figure 15B) from Dr. Bobby Yanagawa, a previous PhD student in our laboratory, showed that the expression of contractile and regulatory genes in CVB3-infected mouse hearts is significantly decreased, which is also unexplainable by dystrophin cleavage.  4.3  Hypothesis and specific aims The objective of the present chapter is to explore viral protease substrate that contributes  significantly to the impaired heart function and development of DCM. Specifically I HYPOTHESIZE that the serum response factor is cleaved by viral protease 2A during CVB3 infection and contributes to impaired myocardial function and progression to DCM. My SPECIFIC AIMS are: Aim 1. Examine whether SRF is cleaved in CVB3 infection Aim 2. Determine the mechanisms of SRF cleavage Aim 3. Determine the functional significance of SRF cleavage  53  4.4  Results  4.4.1  Cardiac function and cardiac gene expression following CVB3 infection The mouse model of myocarditis has been well characterized and commonly used to study  target organ injury during CVB3 infection (34, 133). Two-dimensional echocardiography was performed to assess functional alterations in CVB3-infected mouse hearts. As shown in Figure 15A, virus-infected mice displayed cardiac dysfunction, evidenced by a significant reduction in left ventricular posterior wall (LVPW) thickening and ejection fraction. Heart failure has been associated with the reduction of a broad range of cardiac-specific genes (17, 90, 187). To confirm the downregulation of contractile and regulatory genes in CVB3-infected mouse heart (as shown in Affymetrix microarray data Figure 15B), I performed real-time RT-PCR examination of CVB3-infected cardiomyocytes. Quantitative RT-PCR results in Figure 15C demonstrate downregulation of several cardiac genes, including ANP, β-MHC, GATA-4, MEF2, and NFAT, in CVB3infected cardiomyocytes.  54  P<0.05  A  P<0.005 30  70  % Changes in ejection f raction (As compared to day 0)  LVPW thickening (%)  80  60 50 40 30 20 10 0  0  3  9  Sham  30  0  3  9  30  day pi  20 10 0 -10  Sham CVB3  -20  P<0.01  -30  3  CVB3  9  30  day  pi  C  B  Fold change (log2) (compared to sham)  -0.5 -1 -1.5 -2 -2.5 -3 -3.5  Fold change (compared to sham)  0.0 0  -2.0 -4.0 -6.0 -8.0 -10.0 -12.0  -4  Figure 15. Cardiac function and cardiac gene expression following CVB3 infection. (A) Cardiac function following CVB3 infection by echocardiography. At day 0 (before infection), and 3, 9, and 30 days following CVB3 or sham infection of A/J mice, systolic and diastolic parasternal long and short axis measurements were obtained. Left ventricular posterior wall (LVPW) thickening and ejection fraction are presented as percentage changes (mean ± SD, n = 4 for each group). pi, post infection. (B) CVB3- or sham-infected mouse hearts (n = 4 for each group, pooled) at 9 days pi were collected for Affymetrix array analysis. Gene changes in intensity (averages of two replicates of the microarray analysis) were plotted as a ratio of CVB3- to sham-infected hearts for visualization. Only genes that exhibited a 1.8-fold change (log2) or greater were included. MHC, myosin heavy chain; MLC, myosin light chain. (C) Murine HL-1 cardiomyocytes were sham- or CVB3-infected for 24 h. Real-time quantitative RTPCR was performed to examine the expression of indicated genes. The gene expression was normalized to GAPDH mRNA, and then displayed as fold changes compared to sham infection (mean ± SD, n = 3). ANP, atrial natriuretic peptide; MEF2, myocyte enhancer factor 2; NFATc, nuclear factor of activated T cells.  55  4.4.2  Expression levels of SRF following CVB3 infection As alluded to earlier, SRF is a cardiac-enriched transcription factor, regulating the expression of  numerous cardiac-specific genes. To explore whether downregulation of cardiac genes is associated with dysregulation of SRF, I examined the expression levels of SRF during CVB3 infection. Using an antibody against the C-terminus of SRF, I found that CVB3 infection of mouse cardiomyocytes (Figure 16A, top panel) and hearts (Figure 16D) led to marked decreases in the protein expression of SRF (~67 kDa), accompanied by the appearance of ~20 kDa fragments. To validate the results, I used an antibody against the N-terminus of SRF (Novus Biologicals). As shown in Figure 16A bottom panel, an extra band at the molecular weight of ~50 kDa was detected in addition to the full-length SRF following CVB3 infection of mouse cardiomyocytes at 20 h and 24 h. These data suggest that SRF is potentially cleaved during CVB3 infection to generate two cleavage products of approximately 50 and 20 kDa, respectively. I further examined the distribution of SRF following CVB3 infection of mouse cardiomyocytes using anti-Cterminal SRF antibody. Figure 16B showed that SRF was redistributed from the nucleus to the cytoplasm in CVB3-infected cells, whereas it remained in the nucleus of sham-infected cells. Real-time RT-PCR was also carried out to examine the gene expression of SRF after CVB3 infection of mouse cardiomyocytes. The results in Figure 16C demonstrate unchanged mRNA levels of SRF following CVB3 infection, suggesting that the reduced protein expression of full-length SRF is unlikely due to decreased gene expression, but rather protein cleavage.  56  A  B SRF  DAPI  MW (kDa)  VP1  Overlay  CVB3 (pi) 16h  20h  24h  SRF (short exposure)  64  SRF (long exposure)  49  SRF isof orm?  Sham  Sham 8h  64  37  26  VP1 β-actin  CVB3 (pi) Sham 8h  16h  20h  24h  C SRF Cleaved SRF  64 49  VP1 β-actin  Relative SRF gene expression (SRF/GAPDH)  MW (kDa)  CVB3  Cleaved SRF  19  D Relative SRF protein levels (SRF/GAPDH)  P<0.005  1.0  P<0.005  MW (kDa)  1.8 1.6 1.4 1.2 1 0.8 0.6 0.4 0.2 0  Sham  Sham  64  CVB3  CVB3 (9d pi)  SRF SRF isof orm?  49  0.8  37  0.6 0.4  26  0.2  19  0.0 Sham 1 (n=4)  CVB3,23d pi CVB3,3 9d pi (n=5) (n=4)  Cleaved SRF GAPDH  Figure 16. Protein and gene expression of SRF following CVB3 infection. (A) Protein expression of SRF following CVB3 infection of murine HL-1 cardiomyocytes. HL-1 cells were either sham infected or infected with CVB3 for various times as indicated. Western blotting was performed to examine protein expression of SRF (top, using anti-C-terminal SRF antibody; bottom, using anti-N-terminal SRF antibody), viral protein VP1 and β-actin (loading control). (B) Protein distribution of SRF following CVB3 infection of HL-1 cells. HL-1 cells were infected with CVB3 for 20 h, doubleimmunocytochemical staining was then performed using anti-C-terminal SRF antibody and anti-VP1  57  antibody to examine the expression and localization of SRF (red) and viral protein VP1 (green), respectively. The nucleus was counterstained with DAPI (blue). (C) Gene expression of SRF after CVB3 infection. RNA samples were extracted from HL-1 cardiomyocytes at 24 h following CVB3 infection. Gene levels of SRF were measured by real-time quantitative RT-PCR and normalized to GAPDH mRNA (mean ± SD, n = 3). (D) SRF expression in A/J mouse heart following 3 and 9 days of CVB3 infection. Heart extracts were used for western blot analysis of protein expression of SRF (using an anti-C-terminal SRF antibody) and GAPDH (loading control). Left panels, levels of SRF were quantitated by densitometric analysis using NIH ImageJ V1.43, normalized to GAPDH, and presented as mean ± SD. Right panels, representative western blot of SRF expression in mouse heart at 9 days post infection.  58  4.4.3  Cleavage of SRF following CVB3 infection To further confirm cleavage of SRF, I utilized a HeLa cell line, a well-characterized model in CVB3  investigations amenable for transfection with plasmids. HeLa cells which express low levels of endogenous SRF were transiently transfected with a 3×FLAG-tagged SRF construct for 48 h followed by CVB3 infection for 7 h. Western blot was performed using anti-FLAG antibody or anti-C-terminal SRF antibody, which showed two cleavage products with molecular weights of ~50 and ~20 kDa, respectively (Figure 17). These results correspond well with my observation on endogenous SRF cleavage shown in Figure 16A.  59  A  SRF  Anti-FLAG Antibody  Cleavage  Anti-SRF Antibody  1  508  3xFLAG  B  Exon1  MW (kDa)  + -  + + -  Exon3  Exon2  + -  + + -  + +  CVB3 3 FLAG-SRF Vector control  ×  Exon4  Exon5  Exon6  Exon7  COOH  Long exposure  60  SRF  50  Cleaved SRF VP1 β-actin  C  MW (kDa)  + -  + + -  + -  + + -  + +  CVB3 3 FLAG-SRF Vector control  ×  Long exposure SRF  60  NS 20  Cleaved SRF VP1 β-actin  Figure 17. Cleavage of SRF following CVB3 infection. HeLa cells were transiently transfected with a plasmid expressing 3×FLAG-SRF for 48 h, followed by CVB3 infection for 7 h. (A) Schematic diagram of the 3×FLAG-SRF construct and western blot detection of SRF with the antibodies described in B and C. Western blotting was performed to examine protein expression of viral protein VP1, β-actin (loading control), and SRF using an anti-FLAG antibody that recognizes the N-terminus of SRF (B) or using an anti-C-terminal SRF antibody (C). NS, nonspecific bands.  60  Caspase-mediated SRF cleavage, which yields a 32-kDa protein fragment, has been reported in both non-cardiac and cardiac cells (21, 32, 53). To determine whether caspases are involved in CVB3induced cleavage of SRF, I treated cells with z-VAD-fmk, a general caspase inhibitor. As shown in Figure 18, following CVB3 infection of SRF-transfected cells (lane 6), two cleavage products (~50 and ~40 kDa using anti-N-terminal FLAG antibody and of ~30 and ~20 kDa using anti-C-terminal SRF antibody) were detected. With addition of z-VAD-fmk (lane 7), the levels of the ~40 kDa (upper panel) and the ~30 kDa (middle panel) fragments were markedly reduced, indicating that they are the cleavage products of SRF by caspases. The appearances of the ~40 kDa (upper panel) and the ~30 kDa (middle panel) bands in SRF-transfected and sham-infected cells (lane 5) are likely due to transfection-induced cytotoxicity. It was noted that inhibition of caspase activation did not prevent the production of the ~50 kDa (lane 7, upper panel) and the ~20 kDa (lane 7, middle panel) fragments, suggesting that these cleavages are independent of caspase activation. Taken together, my results suggest that SRF is cleaved during CVB3 infection by viral protein and caspases at two different sites. Since caspase activation is a late event during viral infection, downregulation of cardiac genes in CVB3-infected mouse heart and cardiomyocytes may be mainly attributable to viral protease-mediated SRF cleavage.  61  Figure 18.. Effect of caspase inhibition on CVB3 CVB3-induced cleavage of SRF. HeLa cells were transiently transfected with a plasmid expressing 3×FLAG-SRF SRF or an empty vector for 48 h, followed by CVB3 infection for 7 h in the presence or absence of zz-VAD-fmk fmk (50 μM), a pan caspase inhibitor. Western blotting was performed to examine protein expression of SRF (top, using anti-FLAG anti antibody; middle, using anti-C-terminal terminal SRF antibody), viral protein VP1, and β β-actin actin (loading control).  62  4.4.4  Identification of SRF cleavage site during CVB3 infection As illustrated in Figure 19A, SRF protein consists of two major domains: the SRE DNA binding  and dimerization domain (amino acids 133 to 222) and the transactivation domain (amino acids 339 to 508). Computer prediction algorithm for cleavage sites by enteroviral proteases (NetPicoRNA V1.0 algorithm) revealed potential 2A protease site, which results in two products of ~50 and ~20 kDa (Figure 19A), consistent with the SRF fragments detected in CVB3-infected cells. First, I examined whether SRF can be cleaved by viral protease 2A. Figures 19B and 19C showed that overexpression of 2A resulted in two cleavage products of FLAG-tagged SRF with the molecular weights similar to those observed during CVB3 infection. This study suggests that CVB3 induces SRF cleavage through the action of viral protease 2A.  63  A  SRF  N  Pos  Clv*  Surf#  G  348  0.582  0.712  FTLMPGGAVAQ  Mouse, Rat  G  344  0.582  0.712  FTLMPGGAVAQ  DNA binding and dimerization domain (MADS box---133-222)  Exon1  Exon2  Exon3  C  B MW (kDa)  60  Sequence  Human  Nuclear localization signal (95-100)  1  Residue  Exon4  Transcriptional activation domain (339-508) 508 Exon5  MW Control Control 2A (kDa) 60  Exon6  Exon7  aa C  2A SRF  Control 2A Control 2A SRF NS  50  Cleaved SRF eIF4γ Cleaved eIF4γ β-actin  20  Cleaved SRF  eIF4γ Cleaved eIF4γ β-actin  Figure 19. Cleavage of SRF by overexpressing viral protease 2A. (A) SRF prediction cleavage site for viral protease 2A. The SRF protein sequence was searched for putative cleavage sites of picornavirus proteases using NetPicoRNA V1.0 algorithm. *Cleavage prediction score above 0.500 are predicted as potential cleavage sites. #Surface exposure scores above 0.500 are predicted as surface exposed sites. (B, C) HeLa cells were transiently transfected with 3×FLAG-tagged SRF construct for 48 h, followed by additional transfection with either empty vector (control) or viral protease 2A plasmid for another 48 h. Western blotting was performed to examine protein expression of SRF using anti-FLAG antibody (B) that recognizes the N-terminus of SRF, or by anti-C-terminal SRF antibody (C). The 2A proteolytic activity was confirmed by the cleavage of eIF4γ. Expression of β-actin was shown as a loading control. NS, nonspecific bands.  64  To confirm the cleavage site predicted by the computer algorithm, I constructed a SRF point mutant, in which the G residue at position 349 was replaced with E (G349E), by site-directed mutagenesis. However, I found that this mutant was still susceptible to cleavage during infection (Figure 20B, upper). To further identify the cleavage site of SRF, I constructed three deletion mutants and a series of point mutation plasmids (data not shown). I found that the G327E mutant was resistant to CVB3-induced cleavage (Figure 20B, lower), suggesting that SRF is cleaved after T326. Although this cleavage site is not predicted by the NetPicoRNA program, amino acid sequence alignments of SRF with other known 2A substrates at the cleavage region revealed a similar cleavage motif (Figure 20C).  65  A  327  SRF  349  N -……VLKSTGSGPVSSGGLMQLPTSFTLMPGGA……- C G327E  G349E  B  C  MW (kDa)  Control  SRF-WT  SRF-G349E  P4 P2  Sham CVB3 Sham CVB3 Sham CVB3  60  CVB1  TRSNITTT  GAFGQQSG  CVB2  KRDSLTTT  GAFGQQSG  CVB3  SITTMTNT  GAFGQQSG  CVB4  ERASLITT  GPYGHQSG  CVB5  EITAMQTT  GVLGQQTG  Murine Dystrophin  RAPGLSTT  GASASQTV  SRF  Human Dystrophin  LAPGLTTI  GASPTQTV  Cleaved SRF  Human eIF4γ  GRTTLSTR  GPPRGGPG  VP1  Rabbit eIF4γ  GRPALSSR  GPPRGGPG  β-actin  SRF  NGTVLKST  GSGPVSSG  Cleaved SRF  50  VP1 β-actin  MW (kDa) 60 50  P1’  SRF  Control  SRF-WT  SRF-G327E  Sham CVB3 Sham CVB3 Sham CVB3  Cleavage site  Figure 20. Cleavage of SRF after T326 following CVB3 infection. (A) Diagram of point mutation constructs of SRF. (B) HeLa cells were transiently transfected with 3×FLAG-tagged SRF-wild-type (SRF-WT) or SRF mutants (SRF-G349E or SRF-G327E as indicated) for 48 h, followed by 7 h CVB3 infection. Transfection of an empty vector (pcDNA) was used as negative controls. Western blotting was performed to examine protein expression of SRF using anti-FLAG antibody and viral protein VP1. Expression of β-actin was shown as a loading control. . (C) Identification of protease 2A substrate cleavage site. Substrate recognition by 2A depends on a degenerate amino acid pattern upstream of the cleavage site. The cleavage recognition site usually contains a T, or S at position P2 and an L, I, or M at position P4. A glycine residue at the P1’ C-terminal side of the scissile bond of the cleavage site occurs in all known substrates of the 2A protease.  66  4.4.5  Cellular localization and transcriptional function of CVB3-mediated SRF cleavage  fragments To understand the functional significance of CVB3-induced SRF cleavage, I generated two constructs expressing the N- and C-terminal SRF fragments, respectively. I showed that the SRF-N truncate harboring a DNA-binding domain and a nuclear localization signal was localized to the nucleus in both HeLa cells (Figure 21B) and HL-1 cardiomyocytes (Figure 21C) as expected, whereas SRF-C which carries a transcriptional activation domain was distributed in the cytoplasm (Figure 21B) or in both cytosol and nucleus (Figure 21C). To determine the transcriptional activities of cleaved or non-cleavable SRFs, HeLa cells (Figure 21D) and HL-1 cardiomyocytes (Figure 21E) were transiently co-transfected with Firefly luciferase reporter construct containing cardiac α-actin promoter together with plasmids encoding full-length, truncated, or non-cleavable SRFs and LacZ (control). As displayed in Figure 21D and 21E, the cardiac αactin promoter activities were strikingly increased with the addition of wild-type and non-cleavable forms of SRF. However, both SRF-N and SRF-C mutants lost their gene transcriptional functions. My results suggest that CVB3-induced cleavage of SRF leads to the disruption of SRF-mediated gene transactivation. In addition to loss of function of SRF, the presence of SRF cleavage products may themselves have detrimental effects on the function of native SRF. The N-terminal fragment of SRF contains an intact DNA-binding domain but lacks the C-terminal transactivation domain. To test whether this fragment can act as a dominant-negative competitor to full-length SRF, I co-transfected wild-type SRF together with SRF-N and SRF-C mutants in the presence of cardiac α-actin reporter and Renilla luciferase control plasmids into HeLa cells (Figure 21D) and HL-1 cardiomyocytes (Figure 21E). I demonstrated that the SRF-N mutant significantly decreased wild-type SRF-induced promoter activities of cardiac α-actin, suggesting a negative regulatory role of this mutant in regulating SRF function probably by competing for DNA binding.  67  A  DNA binding/dimerization NLS domain (MADS box)  SRF-WT  Transcriptional activation domain  3xFLAG  C 1  SRF-N  95 100 133  222  339  3xFLAG  C 1  327 aa  SRF-C  3xFLAG  C 328  SRF-G327E  508 aa  3xFLAG  C 1  FLAG  B  508 aa  DAPI  G327E  FLAG  Overlay  C  DAPI  508 aa  Overlay  SRF-N  SRF-N  SRF-C  SRF-C  E  D  12  P<0.001  10  P<0.001 P<0.001  10  P<0.001 8  Fold change  Fold change  P<0.001 6  4  8  P<0.005  6 4 2  2  0  0  + LacZ  + SRF-WT  + LacZ  + SRF-WT  68  Figure 21. Cellular localization and transcriptional activity of SRF mutants. (A) Schematic diagram of four 3xFLAG-SRF constructs. NLS, nuclear localization signal. (B, C) HeLa cells (B) or HL-1 cardiomyocytes (C) were transiently transfected with 3×FLAG-tagged SRF-N or SRF-C fragments of the cleaved SRF products for 48 h using Lipofectamine 2000. Immunocytochemical staining for transfected SRF was performed using anti-FLAG antibody. Nucleus was counterstained with DAPI. (D, E) HeLa cells (D) or HL-1 cardiomyocytes (E) were transiently co-transfected with cardiac α-actin luciferase reporter plasmid and constructs expressing SRF-WT or empty vector and/or SRF-G327E, SRFN, and SRF-C mutants as indicated for 48 h. Luciferase assay was performed and the values are presented as folder changes (mean ± SD, n = 3) over those of control reporter (LacZ), which were arbitrarily set as 1.  69  4.4.6  Effects of CVB3-mediated SRF cleavage on viral replication Viruses usually utilize or modulate the host cell infrastructure for their own replication. To test  whether cleavage of SRF is beneficial to viral propagation within the host cells, finally, I examined the influences of SRF cleavage on viral replication. Figure 22 showed that neither knockdown of SRF nor overexpression of wild-type or truncated SRFs affects viral replication, suggesting that cleavage of SRF may not contribute directly to viral benefits.  A  B Sham  CVB3  siCON siSRF siCON siSRF VP1  Control SRF-N  Control SRF-WT  SRF β-actin  VP1  VP1  SRF  SRF  β-actin  β-actin  CVB3 titer (×106 pfu/ml)  12 10  Control SRF-C  8  VP1  6 SRF  4  β-actin  2 0  siCON  siSRF  Figure 22. Effects of SRF dysregulation on CVB3 replication. (A) Effects of knockdown of SRF on CVB3 replication. The gene expression of SRF was knocked down by siRNA. siCON, scramble siRNA control. Upper, western blot analysis was carried out to examine protein level of viral protein VP1, SRF (with anti-FLAG antibody), and β-actin (loading control). Lower, plaque assay was performed to examine virus titers. The results are shown as mean ± SD (n = 3). (B) Effect of overexpression of SRF-wild-type (SRF-WT) and two truncated SRF mutants on CVB3 replication. HeLa cells were transiently transfected with SRF-WT or truncated SRF mutants (SRF-N or SRF-C as indicated) for 48 h, followed by 7 h CVB3 infection. Western blot analysis was carried out to examine protein level of viral protein VP1, SRF (with anti-FLAG antibody) and β-actin (loading control).  70  4.4.7  Attenuation of SRF cleavage using peptide-based inhibitor Blocking SRF cleavage is a potential therapeutic strategy in the treatment of viral myocarditis.  Viral protease 2A has a catalytic cysteine residue and its substrate recognition is dependent on the 4~6 amino acids upstream of the scissile bond (196). A 2A substrate analogue inhibitor (z-LKST-fmk, shown in Figure 23A) was synthesized based on the identified cleavage site of SRF using a strategy described previously (21), which is similar to that used to inhibit caspases (z-VAD-fmk). Fluoromethyl ketone (fmk) can form a covalent bond with the catalytic cysteine leading to protease inactivation. The effectiveness of z-LKST-fmk in blocking SRF cleavage was evaluated. As shown in Figure 23B, z-LKST-fmk attenuated CVB3-induced SRF cleavage in a dose dependent manner, which was not observed in the z-VAD-fmk treated murine cardiomyocytes.  A SRF:  327 ……VLKSTGSGPVSSGG…… z-LKST-fmk  cbz-leucine-lysine-serine-threonine-fluoromethyl ketone  Chemical Formula: C28H51FN5O8 Molecular Weight: 604.73  71  B CVB3 Z-LKST-fmk (µ µ M)  Sham 0  0  6.25  12.5  25  50  75 SRF  VP1  β-actin Sham  CVB3 O/N  Figure 23. Attenuation of SRF cleavage using peptide-based inhibitor. (A) Design and structure of 2A substrate analogue inhibitor (z-LKST-fmk). (B) Western blot analysis of SRF, VP1, and β-actin (loading control) expression in CVB3-infected HL-1 murine cardiomyocytes treated various dosage of SRF-based 2A inhibitor (z-LKST-fmk) as indicated, or general caspase inhibitor z-VADfmk (50 µM).  72  4.5  Discussion Enteroviral proteases have been recognized as an important contributor to viral pathogenesis  and resultant myocardial dysfunction via the cleavage of important host proteins. Using a Cre/LoxPinducible system, it has been shown that transgenic mice with cardiac-specific overexpression of protease 2A develop severe cardiac dysfunction and DCM (205). Experimental evidence from cell culture and cell-free in vitro studies demonstrates that viral protease 2A induces the cleavage and functional impairment of dystrophin (11-13). Dystrophin connects the cytoskeletal actin-binding site to the βdystroglycan extracellular matrix anchor, thus its cleavage leads to the disruption of the cytoskeletal architecture. It is therefore proposed that protease 2A-induced DCM is associated with disrupted myocyte integrity via the cleavage of dystrophin (13). However, the mdx (dystrophin deficient) mice display a relatively mild dilated phenotype. This is attributed to the compensatory upregulation of the dystrophin homolog utrophin, as evidenced by severe dystrophic phenotype in mice with double knockout of utrophin and dystrophin (50, 76, 80, 93). Previous study has shown that CVB3 infection does not lead to cleavage of utrophin (13), thus the DCM phenotype induced by dystrophin cleavage may be dampened by the compensatory effect of utrophin in the CVB3-infected heart. All of these suggest that dystrophin cleavage alone may not be sufficient to explain the severe cardiomyopathy phenotype observed in 2A-transgenic mice. This prompts us to postulate that other substrate(s) of 2A may also play a role in the progression of virus-induced cardiomyopathy. The major findings of this study are as follows: (1) cardiac-specific contractile and regulatory genes are downregulated in CVB3-infected mouse heart and cardiomyocytes, (2) SRF is cleaved by viral protease 2A during CVB3 infection, and (3) cleavage of SRF leads to the disruption of SRF-mediated gene transactivation and production of a dominant-negative competitor to native SRF. These findings suggest that SRF cleavage may contribute to cardiac dysfunction and subsequent progression to DCM in enteroviral myocarditis by disturbing the expression of cardiac contractile and regulatory genes. Microarray studies in human end-stage heart failure revealed a significant downregulation of numerous cardiac genes, including genes with SRF-binding sites, implicating an important role of SRF in the progression of heart failure (17, 90, 187). Furthermore, as illustrated in this chapter, many cardiacspecific genes are also downregulated in enterovirus-infected heart and cultured cardiomyocytes. All these genes shown in Figure 15 have either been previously reported to be targets of SRF or contain SREs in their promoter regions (14, 31, 183).  73  It was reported that expression of an alternatively spliced SRF variant lacking portions of the transactivation domain is markedly increased in human and animal failing hearts (20, 46). This isoform functions as a dominant-negative mutant that inhibits SRF-dependent activation of cardiac muscle genes. Interestingly, my collaborator’s recent report showed that SRF cleavage also occurs in myocytes of severely failing human hearts. Increased caspase activity during heart failure induces the cleavage of SRF, producing a truncated protein that lacks the C-terminal transactivation domain, and acts as a dominant inhibitory transcription factor, similar to the alternatively spliced variant (32). These studies suggest that decreased production of full-length SRF, or increased accumulation of truncated SRF, may downregulate SRF-dependent genes and contribute to the progression of severe heart failure. In this study, I have identified a new cleavage site in SRF by viral protease 2A during virus infection and demonstrated a dominant-negative effect of the cleaved SRF on the expression of cardiac genes. With this knowledge, I developed a peptide-based SRF cleavage inhibitor (z-LKST-fmk) that effectively attenuated CVB3-induced SRF cleavage in murine cardiomyocytes. Future studies are required to investigate the therapeutic potential of the peptide-based inhibitor. Coxsackievirus infection of heart results in focal lesions accompanied by severe fibrosis. Interestingly, recent research on mosaic regional inactivation of SRF in mouse hearts was found to be sufficient in inducing focal lesions and resulting in fatal DCM (69). In these mice, wild-type cardiomyocytes adjacent to the SRF-null myocytes undergo compensatory hypertrophy, which further triggers a hypertrophic response in remote wild-type cardiomyocytes to preserve cardiac output. This mosaic SRF inactivation model provides some insights into the possible pathophysiological consequences of non-functional SRF and dominant-negative SRF truncates presented in patches of CVB3-infected cardiomyocytes and their subsequent influences on the disease progression and outcome. Overexpression of SRF has also been associated with cardiac hypertrophy. Transgenic mouse studies showed that the cardiac-specific overexpression of SRF induces a pattern of fetal gene reexpression similar to that observed in cardiac hypertrophy and subsequently causes a severe hypertrophic phenotype (212). Therefore, in the future, establishment of a knockin mouse model expressing non-cleavable SRF in the heart will be crucial to determine whether this form of SRF can rescue cardiomyocytes from CVB3-induced dysfunction.  74  4.6  Limitations and solutions In this study I tried to apply multiple experimental methods to ensure the validity of each  finding. For example, I confirmed CVB3-induced SRF cleavage and the expression of its cleavage products in three different models (mouse hearts, murine cardiomyocytes, and HeLa cells expressing exogenous SRF), as well as using different antibodies against SRF to ensure the validity of western blot results. I differentiated viral protease 2A-induced SRF cleavage from caspase 3-induced SRF cleavage using the pan caspase inhibitor z-VAD-fmk. To assess the consequence of SRF cleavage, I studied the intracellular localization and the transcription activation function of SRF cleavage fragments. However, several supplemental experiments could further strengthen my observations. In particular, my data demonstrating viral protease 2A expression is sufficient in inducing SRF cleavage can be supported by further experiments. The expression of SRF cleavage products is low in HeLa cells transfected with viral protease 2A constructs (Figure 19), which is attributed to the inherent limitation of the 2A construct. Protein expression of viral protease from this construct requires the cap-dependent mRNA translation machinery. As described previously, protease 2A effectively shuts down the cap-dependent translation pathway by cleaving eIF4G and PABP. As a result, existing 2A will terminate further expression of 2A. This may be circumvented by constructing an internal ribosome entry site (IRES)-regulated 2A construct, which directs protease 2A expression via the cap-independent translation pathway. Alternatively, an in vitro cleavage assay involving purified viral protease 2A and SRF will help confirm 2A-induced SRF cleavage. Additional specificity can be achieved with the addition of cleavage inhibitor z-LKST-fmk. Regarding to the functional consequences of CVB3-induced SRF cleavage, several supplementary experiments will help strengthen my finding. An electrophoretic mobility shift assay (EMSA) can be conducted to examine the DNA binding activity of truncated SRFs. Other experiments for assessing the functional consequence of SRF cleavage include measuring cardiomyocytes contractility and cardiac gene expression in myocytes transfected with adenovirus constructs overexpressing truncated or noncleavable SRFs. Intracellular localization difference of SRF-C in HeLa cells and HL-1 cells (Figure 21B and 21C) implies different SRF activity in these cell lines. SRF-C, which lacks nuclear localization signal, should reside predominantly in the cytosol as observed in HeLa cells (Figure 21B). However, SRF-C is present evenly in both nucleus and cytosol in HL-1 cells (Figure 21C). As previously described, SRF interacts with its cell-specific cofactors to control gene expression in a cell- and context-specific manner. It is likely certain muscle specific cofactors present in HL-1 cells may still interact with the truncated SRF-C and  75  assist its nuclear shuttling. In this regard, further experiments should be performed to confirm the findings of the effects of CVB3-mediated SRF cleavage on viral replication. HeLa cells were used in this set of experiments to achieve high transfection efficiency. However, HeLa cells are less dependent on SRF-mediated gene expression as reflected from their low endogenous expression level of SRF. Similarly, the expression profile of SRF cofactors is also different in HeLa cells as compared to cardiomyocytes. Therefore, it is important to repeat the same SRF knockdown and overexpression study in cardiac cells such as rat neonatal cardiomyocytes and HL-1 cells to verify the effect of SRF cleavage on CVB3 replication.  4.7  Future directions This study demonstrated a novel viral mechanism in the downregulation of cardiac gene  expression in CVB3-infected host. Coxsackieviral protease 2A-induced SRF cleavage represses cardiac gene expression through the loss of full-length SRF proteins, as well as the generation of dominant negative SRF-N. Future research is required to clarify the role of CVB3-induced SRF cleavage in the progression of viral myocarditis using animal models. The significance of dominant negative SRF-N in the development of viral-induced DCM can be investigated by establishing a transgenic mouse model with cardiac-restricted expression of SRF-N. It answers the question whether SRF-N alone is sufficient to induce cardiomyopathy. Furthermore, the significance of SRF cleavage in the overall development of viral myocarditis can be examined by generating a non-cleavable-SRF-mutant knockin transgenic mouse model. Specifically, a knockin non-cleavable-SRF-mutant mouse model, i.e. homologous replacement of endogenous SRF alleles, is necessary because SRF expression must be strictly controlled in order to maintain cardiac homeostasis. Either overexpression or knockdown manipulation of SRF gene expression in mouse heart results in cardiomyopathy. Based on the identified SRF cleavage site, a peptide-based 2A inhibitor (z-LKST-fmk) was synthesized. This inhibitor effectively attenuated CVB3-induced SRF cleavage in murine cardiomyocytes. The therapeutic potential of this inhibitor in the treatment of CVB3-induced myocarditis warrants further evaluation. Various cell-free assays (e.g. in vitro SRF cleavage assays, as well as caspase, calpain, and proteasome activity assays) will help validate the efficacy and specificity of z-LKST-fmk in blocking protease 2A-induced SRF cleavage. Moreover, functional assessments, such as gene expression and contractility assays, will help evaluate the effectiveness of z-LKST-fmk in preserving the viability of CVB3infected cardiomyocytes. Eventually, in vivo application of z-LKST-fmk will help examine its therapeutic potential in the overall development of coxsackieviral myocarditis/DCM.  76  4.8  Summary I demonstrated that CVB3 manipulates SRF expression via protein cleavage (summarized in  Figure 24). SRF is a transcription factor vital for the expression of cardiac contractile and regulator genes, as well as gene silencing microRNAs. Cardiac-specific knockout of SRF in adult transgenic mice results in disruption of cardiac gene expression and development of severe DCM. I showed that SRF is cleaved in CVB3-infected mouse hearts and cardiomyocytes. Further studies revealed that SRF is cleaved at the 327 amino acids position by CVB3-encoded protease 2A. I demonstrated that SRF cleavage contributes to DCM by abolishing the transactivation property of SRF and generating dominant-negative SRF-truncates. These results provide new insights into the mechanisms by which virus infection impairs heart function and may further the development of new therapeutic approaches to ameliorating myocardial damage and progression to DCM.  77  2A  SRF -C  SRF  SRF -C  SRF  Cleaved SRF fragments SRF -N SRF -N  Transcription  SRE SRF Co-factors  SRF Target Genes Decreased cardiac contractile & regulatory gene expression  Myocyte dysfunction Figure 24. Cleavage of SRF by viral protease 2A leads to myocyte dysfunction. SRF cleavage by viral protease 2A contributes to myocyte dysfunction by the dissociating of the Nterminal DNA binding/dimerization domain from the C-terminal transcriptional activation domain, thus abolishing SRF-mediated gene expression. Furthermore, the N-terminal fragment (SRF-N) exhibits a dominant-negative effect on endogenous SRF function by competing for DNA binding.  78  Chapter 5 – Closing remarks 5.1  Research summary and conclusion In this dissertation, I reported novel host protein manipulation strategies in coxsackievirus-  induced cardiomyopathy. Specifically, my work focused on CVB3’s strategic subversion of the autophagy pathway to support its replication, as well as its cleavage of the serum response factor that contributes to its pathogenicity. A summary of the major findings from my thesis project is listed below: Chapter 3: Autophagy in coxsackievirus infection 1a) CVB3 infection induces the formation of autophagosomes. 1b) Inhibition of autophagy pathway reduces CVB3 replication – Application of autophagy inhibitor 3-methyladenine or siRNA against autophagy pathway components (ATG7, Beclin-1, and Vps34) reduces CVB3 replication. 1c) Increase autophagosome formation enhances CVB3 replication – Autophagy induction by nutrient starvation and treatment with autophagy inducer rapamycin, or Lysosomal inhibition by RNAi against lysosomal protein LAMP2 enhances CVB3 replication. Chapter 4: SRF in coxsackieviral cardiomyopathy 2a) SRF is cleaved in CVB3-infected mouse hearts and murine cardiomyocytes. 2b) SRF is cleaved at the 327 amino acids position by viral protease 2A into two fragments: nuclear N-terminal fragment (SRF-N, ~50 kD) and cytoplasmic C-terminal fragment (SRF-C, ~20 kD). 2c) SRF cleavage contributes to CVB3-induced cardiomyopathy in two folds: 1) SRF cleavage abolishes the transcription activation function of SRF, and 2) it generates the dominant negative SRF-N that inhibits wild-type SRF. Based on these results, I conclude that the subversion of host autophagy protein degradation pathway and the cleavage of serum response factor serve critical roles in the pathogenesis of CVB3induced myocarditis. Strategic subversion of host autophagy machinery is important for efficient viral replication, whereas cleavage of SRF is central to CVB3-induced myocardial dysfunction. Target intervention of these viral strategies may help limit viral replication, ameliorate cardiomyocyte injury, and preserve cardiac function in viral myocarditis.  79  5.2  Research significance The newly discovered viral strategies reported in this dissertation may serve as potential  therapeutic targets for the treatment of viral myocarditis/DCM. As previously described, viral myocarditis is subdivided into three phases: viremic, inflammatory, and remodeling. At viremic phase, where active viral replication takes place, temporal application of autophagy inhibitor may help control viral replication and limit direct viral-induced myocyte damage. As demonstrated in Figure 7, autophagy inhibitor (3-MA) treatment potently inhibited CVB3 replication in vitro, even when introduced at 5 hours post infection. However, the impact of autophagy inhibition beyond the viremic phase requires further investigations. Since the outcome of viral myocarditis also depends on the complex interactions between virus infection and host immune system, the effect of autophagy inhibition on immune response must be considered. Recent studies suggest a role for autophagy in host adaptive immune responses through the delivery of intracellular pathogenic antigens for major histocompatibility complex class II presentation to CD4 T cells (176). In vivo application of autophagy inhibitor will help evaluate its therapeutic potential in viral myocarditis. Autophagy inhibition in the remodeling phase is cautioned since impairment of protein degradation system has been reported in human failing hearts (22, 198). Application of autophagy inhibitor in the later phases of viral myocarditis may further exacerbate myocardial damage by shutting down this alternative protein degradation pathway. As shown in transgenic mice with cardiac-specific ATG5 knockdown, autophagy is essential to cardiac viability (154). These mice developed cardiac hypertrophy, left ventricular dilatation, and contractile dysfunction (154). Therefore, pharmacological interventions of host autophagy pathway must be carefully designed and stringently evaluated to ensure outstanding benefits with minimal side-effects. Viral-induced SRF cleavage represents an important mechanism in cardiac gene downregulation in viral myocarditis/DCM. Blocking SRF-cleavage may be a promising therapeutic strategy in the management of viral myocarditis patients. Based on the catalytic requirement and substrate recognition pattern of viral protease 2A, I designed a SRF-based 2A substrate analogue inhibitor (z-LKST-fmk). Fluoromethyl ketone (fmk) can form a covalent bond with the catalytic cysteine leading to protease inactivation. Preliminary study revealed that z-LKST-fmk attenuates CVB3-induced SRF cleavage in a dose dependent manner. Blocking the cleavage of SRF concurrently eliminated the production of dominant negative SRF-N. 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