A Dissertation Entitled Negative Regulation of Host Innate Immune

A Dissertation

entitled

Negative Regulation of Host Innate Immune Signaling and Response Pathways by Viral

and Host Regulatory Factors.

Qi Ke

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the

Doctor of Philosophy Degree in Biology

______Dr. Douglas W. Leaman, Committee Chair

______Dr. Malathi Krishnamurthy, Committee Member

______Dr. Travis Taylor, Committee Member

______Dr. Carol A. Stepien, Committee Member

______Dr. Scott Leisner, Committee Member

______Dr. Amanda Bryant-Friedrich, Dean College of Graduate Studies

The University of Toledo

This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author. An Abstract of

Negative Regulation of Host Innate Immune Signaling and Response Pathways by Viral and Host Regulatory Factors.

Qi Ke

Submitted to the Graduate Faculty as partial fulfillment of the requirements for the Doctor of Philosophy Degree in Biology

The University of Toledo August 2016

Eukaryotes have evolved intricate innate immune systems that allow rapid response to pathogens such as viruses. The initiation of innate immune responses depends on the recognition of pathogen associated molecular patterns (PAMPs) by host germ-line encoded pattern-recognition receptors (PRRs). Activation of antiviral responses by these pathways are intended to slow or contain viral replication until the adaptive immune system can clear the infection.

Viral hemorrhagic septicemia virus (VHSv) is a deadly fish rhabdovirus that infects over 50 species of freshwater and marine fishes around the world. In 2003, a new substrain of VHSv (IVb) was found in the Great Lakes region when it caused a massive die-off of many freshwater species. The VHSv genome is about 11-kb long containing six genes, and replication occurs entirely in the cytoplasm using a combination of virally encoded and host-derived factors. As with other viruses, VHSv must neutralize or evade the host innate immune response in order to survive. We have found that VHSv can inhibit IFN-stimulated antiviral responses. Interestingly, the matrix (M) protein of VHSV

IVb alone can potently suppress MAVS- and IFN-induced gene expression in a dose-

iii dependent manner. The inhibition of constitutive SV40 promoter-driven gene expression by M implicated a general effect on transcription or translation.

Our study showed decreased nascent RNA levels in both VHSv-infected cells and

M-transfected cells. Co-transfection of M with a tetracycline inducible reporter gene

(mouse secreted embryonic alkaline phosphatase - mSEAP) resulted in potent inhibition of tet-induced mSEAP mRNA synthesis. These results suggested that M inhibited protein expression by shutting down host transcription. Indeed, ChIP studies illustrated M- dependent inhibition of RNA polymerase II (RNAP II) recruitment to a gene promoter, and decreased RNAP II CTD Ser2 phosphorylation, an indicator of transcript elongation, during VHSv infection. Therefore we hypothesize that VHSv M inhibits host transcription by preventing RNAP II recruitment or by disrupting its association with target genes. M inhibited pol I, II and III transcription in cell-based luciferase studies, but blocked RNAP II-dependent transcription most potently. When M proteins from a variety of VHSv strains and related fish rhabdoviruses were tested for potency in cell-based luciferase inhibition assays, a VHSv F1 substrain M variant was significantly less potent than M from the IVb substrain. Among the four amino acid differences between the two

M protein, two of them (D62A and E181A) were demonstrated to be crucial for the transcriptional inhibitory effect of M. Reverse genetics studies to introduce these amino acid changes into the VHSv IVb backbone have been initiated to determine how they impact virulence within the context of an intact virus. These studies should enhance our understanding of M’s role in host inhibition and, ultimately, viral replication.

Type I IFNs play important roles in both innate and adaptive immune responses and are highly regulated to limit tissue damage and prevent autoimmunity. A second

iv dissertation project focused on mammalian RNF114, an E3 ubiquitin ligase, which we propose regulates dsRNA-induced antiviral responses. Our work has revealed that the

RNF114 RING finger domain and ubiquitin interacting motif (UIM) are both crucial for its ubiquitination activity. When ectopically expressed, RNF114 negatively impacted cellular dsRNA responsiveness, down-regulated the RLH adaptor molecule MAVS expression, potentially through ubiquitination-dependent degradation, and also suppressed RIG-I, MDA5 and MAVS signaling. In contrast, knocking down RNF114 robustly promoted cellular dsRNA responses. RNF114 mRNA was expressed in many different tissues in the mouse, but was predominant in spleen. We have successfully generated a RNF114 knockout mouse which exhibited elevated basal and dsRNA induced IFN and ISG56 mRNA levels, as compared to wild type mice. Mouse bone marrow-derived macrophage (BMDM) studies also suggested RNF114 KO leads to augmented mRNA levels of ISG and IL-10 induced by dsRNA treatment. Thus, we hypothesize that RNF114 may function as a regulatory E3 ubiquitin ligase to inhibit uninduced IFN production by regulating MAVS steady state levels through ubiquitination.

Our overall hypothesis is that RNF114 functions to prevent uncontrolled inflammatory signal. Overall, the combined results of our studies emphasize the critical roles of factors within the innate immune response system, as well as external forces that can alter normal innate immune pathway function to the detriment of the host, and reiterate the need for integrated studies from both the host’s and pathogen’s perspective.

Better understanding of type I IFN regulation, in particular, is important to understand a variety of normal, pathogenic and disease states.

v This dissertation is lovingly dedicated to my father, Hongwu Ke and mother, Yaping Liu.

Their support, encouragement, and constant love have sustained me throughout my life.

Acknowledgements

I would like to thank my advisor Dr. Douglas W. Leaman for his patient guidance, generous inspirations and unflagging encouragement. Throughout the years, he was always by my side providing me advice and support, and carrying me through all the frustrations and struggles. I’ve learned a lot from him as a great mentor, a wonderful person, and a passionate scientist. I would also like to thank my PhD committee members:

Dr. Malathi Krishnamurthy, Dr. Travis Taylor, Dr. Scott Leisner and Dr. Carol A.

Stepien, for their time and valuable suggestions, helpful discussions and comments that help me to build my projects. From the department of marine biology at University of

Maryland, I want to thank Dr. Vikram N. Vakharia for his collaboration, support and help.

I want also to thank my present and past lab members: Dr. Boren Lin, Adam Pore, Dr.

Kuladeep Reddy Sudini, Samantha Stefl, Wade Weaver, Shelby Powell, Pham Loc, Dave

Velliquette, Ali Abou-Alaiwi, Sydney Yoho, Emily Scott, Robert Rominski, Daniel

Hettel, Tyler Williams, Tyler Popi, Jacob Blandford, and Dr. Julia Wildschutte for all your help and laughter over these years. I also want to thank all of my friends, especially

Dr. Zheng Ma for her encouragements during my Ph.D. studies. I am very grateful for my father, Hongwu Ke, and mother, Yaping Liu, who inspired me and gave me endless support and love.

vii

Table of Contents

Abstract iii

Acknowledgements vii

Table of Contents viii

List of Tables xi

List of Figures xii

List of Abbreviations xiii

List of Symbols xvi

I. Overall Introduction 1

1.1. General Introduction 1

1.2 Host Innate Immune System 2

1.2.1 RIG-I-like Helicases Pathway 3

1.2.2 Type I IFN and Interferon Stimulated Genes 5

1.3 Virus Evading Host Antiviral Responses 9

1.3.1 Virus Blocking the Innate Immune System 9

1.3.2 Virus Inhibiting Host Antiviral Responses by Other Mechanisms 10

II. Materials and Method 11

2.1 Cell Lines and Culture Conditions 11

viii 2.2 Plasmids 12

2.3 VHSv Stocks and Infection 13

2.4 Cloning and PCR 14

2.5 Transfection 15

2.6 Click-iT Asssay and Immunofluorescence Microscopy 15

2.7 Luciferase Assays/-gal Assays 15

2.8 Immunoblotting 16

2.9 Real-time PCR 17

2.10 Lentivirus 17

2.11 ChIP Assay 18

III. VHSv Matrix Protein Inhibits Host Transcription 21

3.1 Introduction 21

3.1.1 Viral Hemorrhagic Septicemia 21

3.1.2 Viral Hemorrhagic Septicemia Virus 22

3.1.3 Nomenclature and Classification 25

3.1.4 VHSv IVb in the Great Lakes 27

3.1.5 Transmission 27

3.2 Results 29

3.2.1 Vhsv Inhibits Host Antiviral Responses 29

3.2.1.1 Vhsv Ivb M Inhibits Host Gene Expression 29

3.2.2 M Protein Inhibits Host Transcription 36

3.2.2.1 VHSv M Blocks Nascent Cellular RNA Synthesis 36

3.2.2.2 VHSv IVb M Inhibits RNAP I-III 41

ix 3.2.2.3 M Disrupts RNAP II Activity and Recruitment 46

3.2.3 Single Amino-Acid Change in M Affects Its Inhibitory Function 50

3.3 Discussion 55

IV. RNF114 Inhibits RLH-Dependent Antiviral Responses 59

4.1 Introduction 59

4.1.1 Regulation of RLH Pathway by Ubiquitination 59

4.1.2 RNF114 61

4.2 Results 63

4.2.1 RNF114 Negatively Impacts Host Innate Immune Responses 63

4.2.1.1 RNF114 Inhibits Cellular dsRNA Responses 63

4.2.1.2 RNF114 Suppresses RLH-dependent IFNβ Production 69

4.2.2 RNF114 is an E3 Ubiquitin Ligase Targeting MAVS for

Degradation 74

4.2.2.1 RNF114 is an E3 Ubiquitin Ligase 74

4.2.2.2 RNF114 Reduced MAVS Levels 77

4.2.3 RNF114 Inhibits Cellular dsRNA Responses in vivo 81

4.3 Discussion 85

4.4 Overall Summary 89

References 91

List of Tables

Table 1.1 Primers for PCR analysis and cloning...... 19

List of Figures

Fig. 1-1 Virus Detection and Type I IFN Response Pathways...... 8

Fig. 3-1 Schematic of VHSv virion structure and genome...... 23

Fig. 3-2 VHSv-IVb M Inhibits Host Gene Expression...... 33

Fig. 3-3 VHSv M Blocks Nascent Cellular RNA Synthesis...... 38

Fig. 3-4 VHSv-IVb M Blocks Pol I-III Dependent Transcription...... 43

Fig. 3-5 M Disrupts RNAP II Activity and Recruitment...... 48

Fig. 3-6 Single Amino-Acid Change in M Affects its Inhibitory Function...... 52

Fig. 4-1 RNF114 Inhibits Cellular dsRNA Responses...... 65

Fig. 4-2 RNF114 Inhibits RLH-mediated IFN Production...... 70

Fig. 4-3 RNF114 is an E3 Ubiquitin Ligase...... 75

Fig. 4-4 RNF114 Decreased MAVS Levels...... 78

Fig. 4-5 RNF114 Inhibits Cellular dsRNA Responses in vivo...... 82

xii

List of Abbreviations

ATCC ...... American Type Culture Collection

BMDM ...... Bone Marrow-Derived Macrophages

CARD ...... caspase recruitment domain CPE ...... cytopathic effect

DAPI ...... 4',6-diamidino-2-phenylindole DMEM ...... Dulbecco’s Modified Eagle medium DOX ...... doxycycline dsRNA...... double stranded ribonucleic acid

E1 ...... ubiquitin-activating enzyme E2 ...... ubiquitin-conjugating enzyme E3 ...... E3 ubiquitin ligase EPC ...... Epithelioma papulosum cyprinid ER ...... endoplasmic reticulum EU ...... 5-ethynyl uridine

FADD ...... Fas-associated protein with death domain FBS ...... fetal bovine serum

G ...... Glycoprotein GAPDH ...... glyceralde 3 phosphate dehydrogenase

IFITM1 ...... interferon induced transmembrane protein 1 IFN ...... interferon IFNAR...... interferon alpha receptor IHNV...... Infectious hematopoietic necrosis virus IKK ...... IB kinase IRAK2 ...... interleukin 1 receptor-associated kinase 2 IRF ...... interferon regulatory factor

xiii ISG ...... IFN-stimulated gene ISGF3 ...... IFN-stimulated gene factor 3 ISRE ...... IFN-stimulated response element ITS1...... internal transcribed spacer region 1

JAK ...... Janus kinases

L ...... Polymerase protein LGP2 ...... laboratory of genetics and physiology 2 LPS ...... lipopolysaccharide LRR ...... leucine-rich repeat

M ...... Matrix protein MAM...... mitochondiral-associated membrane MAVS ...... Mitochondrial Antiviral Signaling protein MDA5 ...... melanoma differentiation associated factor 5 MEM ...... minimum essential medium MERS ...... Middle East respiratory syndrome

N ...... Nucleoprotein NACHT ...... neuronal apoptosis inhibitory protein, CIITA, HET-E and TP-1 NALP3 ...... neuronal apoptosis inhibitory protein, CIITA, HET-E and TP-1, leucine-rich repeat and pyrin domains-containing protein 3 NEMO ...... nuclear factor kappa-light-chain-enhancer of activated B cells essential modulator (NEMO) NF-B ...... nuclear factor kappa-light-chain-enhancer of activated B cells NV ...... Non-virion protein NLR...... nucleotide-binding oligomerization domain-like receptor NOD ...... nucleotide-binding oligomerization domain NS1 ...... nonstructural protein 1

P ...... Phosphoprotein PAMP ...... pathogen-associated molecular pattern PBS ...... Phospate-buffered saline Poly IC ...... Polyinosinic polycytidylic acid PRR ...... pattern recognition receptor PYD...... pyrin domain

RIG-I ...... retinoic acid-inducible gene-I RING ...... really interesting gene RLR ...... retinoic acid-inducible gene-I-like receptor RVFV ...... Rift Vally fever virus

SARS...... Severe acute respiratory syndrome SARS-CoV ...... SARS-associated coronavirus SHRV ...... Snakehead rhabdovirus

xiv ssRNA ...... single-stranded ribonucleic acid STAT...... signal transducers and activators of transcription SVCV ...... Spring viremia of carp virus

TANK ...... tumor necrosis factor receptor-associated factor member- associated NF-kappa-B activator TBK1...... tumor necrosis factor receptor-associated factor member- associated NF-kappa-B activator-binding kinase 1 TLR ...... Toll-like receptor TRADD ...... tumor necrosis factor receptor superfamily, member 1a-associated via death domain TRAF ...... tumor necrosis factor receptor-associated factor TYK ...... tyrosin kinase

Ub ...... ubiquitin VHS...... Viral hemorrhagic septicemia VHSv...... viral hemorrhagic septicemia virus VSV...... vesicular stomatitis virus VV ...... Vaccinia virus

WHO ...... World Health Organization

List of Symbols

α ...... Alpha β ...... Beta γ ...... Gamma δ ...... Delta ε ...... Epsilon κ...... Kappa λ ...... Lambda ω ...... Omega µ ...... Micro τ ...... Tau

xvi

Chapter 1

Overall Introduction

1.1 General Introduction

Despite advances in medical technology and drug development over the past century, the threat of new emerging or reemerging infectious diseases remains constant.

In 2002, infectious diseases were associated with 26% of human deaths worldwide, representing the second leading cause of death (WHO, 2005). Several recent viral outbreaks have had devastating consequences, especially in tropical or developing countries where medical resources are limited. In February 2003, an epidemic of severe acute respiratory syndrome (SARS) was reported in Asia which caused by a newly identified coronavirus, called SARS-associated coronavirus (SARS-CoV). The disease spread across Asia, Europe, both North and South America infecting 8,098 people and killing 774 before it was contained (WHO, 2005). The Ebola outbreaks in 2014 was the largest in history, infecting more than 27,237 people and killing 11,158 people in West

1 Africa by June, 2015 (WHO, 2015). Recently, another “SARS-like” disease, the Middle

East respiratory syndrome (MERS), which is caused by a novel coronavirus (MERS-

CoV), emerged in the Middle East. The epidemic subsequently spread to Europe, Africa,

North America and Asia, and a severe, ongoing outbreak in Republic of Korea and China has resulted in 16 deaths among 150 confirmed cases reported by WHO (Cheng VC et al.,

2007). MERS-CoV has provoked global concern due to its high fatality rates and limited treatment options. Infectious diseases not only pose a threat to humans, but also have a huge impact on the economy. Avian influenza (AI) viruses, such as A(H5N1) and

A(H7N9), has caused serious human diseases, commonly known as bird flu (WHO,

2004). During the outbreak in 2004, more than 100 million birds either died or were culled in Asia. Countries that are dependent on poultry industries suffered great economic damage. Therefore, understanding viral pathogenesis and host immune responses is very important for drug or vaccine development and prevention of future outbreaks.

1.2 Host Innate Immune System

Higher eukaryotes have evolved complex innate immune systems that serve as the first line of defense against pathogens like bacteria, fungi and viruses. Host cells detect conserved pathogen-associated molecular patterns (PAMPs) via germline-encoded pattern recognition receptors (PRRs; Janeway 1989) and trigger signaling cascades to produce anti-microbial factors such as type I interferons (IFNs) and other cytokines

(Janeway Jr & Medzhitov 2002). Among the best characterized PRRs are the Toll-like receptors (TLRs) that detect lipopeptides, lipoteichoic acid, glucans, proteins, lipopolysaccharide (LPS) and nucleic acids (Akira et al., 2006). TLR1, -2, -4, -5, -6 and -

11 are expressed on the cell surface while TLR3, -7, -8 and -9 are localized in

2 intracellular vesicles such as the endosome or lysosome and the endoplasmic reticulum

(ER). Other classes of PRRs include the cytosolic PRRs such as RIG-I (retinoic acid- inducible gene 1)-like helicases (RLHs) and nucleotide-binding oligomerization domain

(NOD)-like receptors (NLRs). NLR family members such as NOD1 and NOD2 recognize intracellular peptidoglycan-derived bacterial products (Fritz et al., 2006) while other family members promote inflammasome-mediated IL-1/IL-18 maturation in response to a variety of pathogen or cell damage-derived signals (Martinon and Tschopp, 2004, Ogura et al., 2006; Ting et al., 2008).

1.2.1 RIG-I-like Helicases Pathway

The RLHs, RIG-I, MDA5 (melanoma differentiation associated factor 5) and

LGP2 (laboratory of genetics and physiology 2), are cytoplasmic PRRs expressed in both immune and nonimmune cells that are essential for detection of intracellular RNA products, primarily of viral origin (Loo & Gale 2011). Both RIG-I and MDA5 have an N- terminal caspase recruitment domain (CARD) and a DExD/H box RNA helicase domain.

LGP2 lacks the CARD and, although it initially appeared to harbor no signaling capacity, its function in regulating RLHs pathway remains controversial. Several studies suggested that LGP2 inhibits RIG-I activity and can be induced during antiviral responses serving as a feedback inhibitor (Komuro & Horvath, 2006; Rothenfusser et al., 2005; Yoneyama et al., 2005). However, other groups reported LGP2 had a positive role in antiviral signaling pathway. LGP2 deficient mice were more susceptible to viral infection such as

Sendai virus (Suthar et al., 2012). LGP2 was also implicated as a positive regulator of antiviral responses upstream of RIG-I and MDA5 and its ATPase activity is important for the synergistic activation (Satoh et al., 2010). RIG-I recognizes Paramyxoviridae,

3 Rhabdoviridae, Orthomyxoviridae and Flaviviridae members (Kato et al., 2006). RIG-I preferentially binds to viral ssRNA or dsRNA marked with 5’ triphosphate termini

(Hornung et al., 2006; Pichlmair et al., 2006; Yoneyama et al., 2004), dsRNA structure

(Baum et al., 2010), blunt-end (Marques et al., 2006), polyuridine-rich motif (Saito et al.,

2008; Uzri and Gehrke, 2009;), specific processing or cleavage (Malathi et al., 2007,

2010). MDA5 detects Picornaviridae members, murine norovirus (Kato et al., 2006;

McCartney et al., 2008) and high-molecular-weight poly(I:C) fragments (Kato et al.,

2008). In the absence of ligand, the CARD domain of RIG-I is maintained in an inactive state by intramolecular interaction with the repressor domain (RD) within the C-terminal domain (CTD; Kowalinski et al., 2011). Once associated with viral RNA through RIG-I helicase domain and CTD, a conformation change in RIG-I releases the CARD domain from CTD repression (Saito et al., 2007). The exposed CARD domain associates with another CARD containing adaptor protein, MAVS (mitochondrial antiviral signaling protein; also called IPS-1/Cardif/VISA), through CARD-CARD interaction (Kawai et al.,

2005). MAVS is located on the outer mitochondrial membrane and mitochondrial- associated membranes (MAM) of the endoplasmic reticulum and peroxisomes through its

C-terminal transmembrane (TM) domain (Seth et al., 2005; Dixit et al., 2010; Horner et al., 2011). Upon activation, MAVS aggregates by forming filaments that are crucial for recruitment and activation of downstream signaling molecules (Seth et al., 2005; Hou et al., 2011; Tang & Wang, 2009). The proline-rich region of MAVS interacts with tumor necrosis factor receptor-associated factors (TRAFs) family members such as TRAF2,

TRAF3 and TRAF6 (Xu et al., 2005; Saha et al., 2006). TRAF3 activates downstream kinases NF-B activator (TANK)-binding kinase 1 (TBK1) and IB kinase- (IKK)

4 (Häcker et al., 2006). TBK-1 and IKK phosphorylate serine and threonine residue of interferon regulatory factor (IRF) 3 and IRF7 (Yoneyama et al., 1998; Lin et al., 1998;

Fitzgerald et al., 2003) promoting IRF homo- or hetero-dimerization and nuclear translocation, thereby promoting transcription of type I IFNs and other dsRNA/virally regulated genes (Lin et al., 1999). MAVS also associates with TRAF2 and TRAF6 to activate the IKK complex composed of IKK, IKK and NF-B essential modulator

(NEMO), which phosphorylates I-B resulting in its poly-ubiquitination and proteasomal degradation. NF-B is then translocated into the nucleus where it transcriptionally regulates IFNs and inflammatory genes (Akira et al., 2006; Karin & Ben-Neriah, 2000).

1.2.2 Type I IFN and Interferon Stimulated Genes

IFNs were first identified in 1957 and named for their ability to interfere with viral replication (Isaacs & Lindenmann, 1957). IFNs are also involved in regulation of cell growth, angiogenesis and various immunological processes (Stark et al., 1998;

Indraccolo, 2010). IFNs are secreted by cells following exposure to virus and act upon neighboring cells to inhibit viral replication. IFNs are class II cytokines that are further classified into three different types (type I, II and III) based on sequence conservation and evolutionary relationships, as well as cognate receptor complex associations (Levy et al.,

2001; Goodbourn et al., 2000). Type I IFNs are grouped into seven classes: IFN-, IFN-,

IFN- IFN-ɛ, IFN-κ, IFNω and IFN- (Pestka et al., 2004; LaFleur et al., 2001). IFN- is the only member of type II IFN and is mainly secreted by T cells and natural killer cells

(Platanias, 2005). Recently the third type of IFNs was identified, and includes IFN-λ1, -

λ2, and -λ3 (Kotenko et al., 2003; Sheppard et al., 2003). IFN- are the best- characterized type I IFNs and represent a complex gene family, with 13 human and 14

5 murine IFN- subtypes (Hardy et al., 2004). Only one IFN- locus has been identified in human and mice but other species, such as cattle, have multiple loci (Roberts et al., 1998).

Type I IFNs play essential roles in the innate immune system and all share the heteromeric IFN-/ receptor (IFNAR) consisting of IFNAR1 and IFNAR2 subunits

(Kim et al., 1997). IFNβ is expressed in most cell types while IFNα is produced predominantly by leukocytes and dendritic cells (Cella et al., 1999; Siegal et al., 1999).

Once secreted, IFNs associate with the cognate IFNAR complex to alter its conformation and activate associated Janus kinases (JAKs) family members tyrosine kinase 2 (TYK2) and JAK1. Activated JAKs phosphorylate specific tyrosine residues on the cytoplasmic tail of the receptor chains, which recruit signal transducer and activator of transcription-1

(STAT1) and STAT2, which are then also phosphorylated on tyrosine by the JAKs.

Phosphorylated STAT1 (pSTAT1), pSTAT2 and IRF9 form a heterotrimeric complex called IFN-stimulated gene factor 3 (ISGF3), which translocates into the nucleus. ISGF3 binds to IFN-stimulated response elements (ISREs) in the regulatory regions of IFN- stimulated genes to promote their transcription (ISGs; Platanias, 2005). ISG encoded proteins represent components of many cellular functions, including translational regulators, enzymes, proapoptotic and antiapoptotic factors. These factors, numbering at least several hundred, work together to establish an antiviral state (Stark et al., 1998).

The IFN system is highly conserved from mammals down to bony fish and all of the critical signaling molecules in the viral detection and IFN response pathways, including RLHs, JAK and STAT signaling molecules and a number of traditional ISGs such as Mx have been cloned from multiple fish species (Chang et al., 2011; Stafford et al., 2003; Oshiumi et al., 2003; Biacchesi et al., 2009; Zou et al., 2007; Shi et al., 2012;

6 Zhang & Gui, 2004; Verrier et al., 2011). Fish IFNs are similar to mammalian type I

IFNs based on coding sequences and crystal structure analysis, although the fish genes differ from their mammalian counterparts by the inclusion of introns (Hamming et al.,

2011; Qi et al., 2010; Sun et al., 2009; Zou et al., 2007). These studies suggest that the innate antiviral pathways and proteins in fish are likely to share many regulatory features in common with their mammalian orthologs.

Higher Eukaryotic Virus Detection and Type I IFN Response Pathways

IFN Virus Protein Virus dsRNA ssRNA RLHs JAK-STAT T L MAVS mRNA mRNA ISG /IPS1 NF-kB s IRFs IFN Reg. Proteins

Virus detection IFN Response Antiviral Protection

Fig. 1-1. Virus Detection and Type I IFN Response Pathways. The initiation of innate immune responses depends on the recognition of pathogen associated molecular patterns

(PAMPs) by host germ-line encoded pattern-recognition receptors (PRRs). RIG-I-like helicases (RLHs), including RIG-I and MDA5 recognize and associate with dsRNA then transmit the signal to mitochondrial antiviral signaling protein (MAVS, or IPS-1). MAVS recruits downstream signaling molecules to induce type I IFN expression. After synthesis and secretion from the cell, type I IFNs bind to IFN receptors on neighboring cells and activate numerous transcriptional factors to induce expression of interferon stimulated genes (ISGs). ISG proteins, including translational regulators, enzymes, proapoptotic and antiapoptotic factors, work together to establish an antiviral state.

8 1.3 Virus Evading Host Antiviral Responses

1.3.1 Virus Blocking the Innate Immune System

To survive and propagate, viruses must be able to evade or overcome the host immune system (Vossen et al., 2002). As an essential component of the innate immune system, type I IFN production or response pathways are common targets for suppression or evasion by viruses (Goodbourn et al., 2000). The nonstructural protein 1 (NS1) of influenza A virus interacts with RIG-I and MAVS, thereby inhibiting downstream IRF3 activation to suppress IFN production and thus antagonize IFN-induced antiviral responses (Donelan et al., 2003; Mibayashi et al., 2007; Talon et al., 2000). NS1 also inhibits TRIM25α mediated K63-linked ubiquitination of RIG-I preventing RIG-I activation and MAVS binding, again leading shutdown of IFN production (Gack et al.,

2009). The poxvirus Vaccinia virus (VV) protein A52R inhibits TLR-mediated NF-B activation by associating with interleukin 1 receptor-associated kinase 2 (IRAK2) and

TRAF6 thereby disrupting signaling molecule complex formation (Harte et al., 2003).

N1L protein of VV inhibits NF-B and IRF3 activation by Toll/IL-1R signaling by targeting IKK complex, potentially by interacting with TBK1 (DiPerna et al., 2004).

Several viruses such as Respiratory Syncytial virus, Rabies virus and Ebola virus inhibit

IRF3 phosphorylation via inhibition by NS, phosphoprotein or VP35, respectively (Basler et al., 2003; Bossert et al., 2003; Brzózka et al., 2005). V protein of a wide variety of paramyxoviruses interacts with MDA5 via its highly conserved cysteine-rich CTD to inhibit MDA5-stimulated activation of IFN promoter (Andrejeva et al., 2004). These are just a few examples of ways in which viruses antagonize host innate immune responses to allow for replication and dissemination to other hosts.

9 1.3.2 Viruses Inhibit Host Antiviral Responses by Indirect Mechanisms

In addition to specific inhibition of viral detection pathways, viruses can evade host antiviral responses by non-specific repression of IFN, such as inhibiting global gene expression in the host. One example is the rhabdovirus vesicular stomatitis virus (VSV), which potently inhibits host antiviral responses by blocking protein expression via the viral matrix (M) protein (Ahmed & Lyles, 1998). Previous studies suggest that VSV M protein can inhibit both transcription and nuclear-to-cytoplasm RNA export by interacting with the mRNA export factor Rae1 and the nucleoporin Nup98 (Faria et al.,

2005; von Kobbe et al., 2000). Recent studies showed that NSs protein of Rift Vally fever virus (RVFV) inhibits IFNβ expression by blocking cellular transcription (Billecocq et al.,

2004). NSs interacts with the E3 ubiquitin ligase FBXO3 and mediates ubiquitination and proteasomal degradation of the transcription factor TFIIH subunit p62 (Kainulainen et al.,

2014).

Although host innate immune pathways are evolved for protection against pathogens, they are susceptible to inhibition by pathogens via a variety of strategies.

Precise regulation of innate immune system is also critical to maintain immune homeostasis and to avoid chronic inflammation and autoimmunity. In chapter 3 we will assess the impact of VHSv proteins on host responses. The overall goal of this study is to understand how pathogens evade or even manipulate host immune responses in favor of the pathogens and the mechanisms by which these pathways are being regulated in the host.

Chapter 2

Materials and Method

2.1 Cell Lines and Culture Conditions

Epithelioma papulosum cyprinid (EPC) cells were purchased from the American

Type Culture Collection (ATCC) (Rockville, MD). The cells were grown in minimum essential medium (MEM) (Fisher) supplemented with 10% fetal bovine serum (FBS)

(Invitrogen) and 1% penicillin/streptomycin (Invitrogen) at 20° C in a 5% CO2 enriched environment. HT1080, FEMX, A375, HEK-293, L929 cells were cultured in Dulbecco’s

Modified Eagle medium (DMEM) (Fisher) supplemented with 10% FBS and 1% penicillin/streptomycin at 37° C in a 5% CO2 enriched environment. To generate

RNF114 shRNA stable knockdown FEMX cell lines, FEMX cells were infected with lentiviruses carrying shRNA against RNF114 (AddGene) for 4 days, and then treated with 1 g/ml Puromycin (Sigma-Aldrich, MO) to select puromycin resistant clones.

Individual clones resistant to puromycin were selected and screened for RNF114

11 knockdown. Bone Marrow-Derived Macrophages (BMDM) were developed by using standard, published protocols (Cold Spring Harb. Protoc., 2008). Briefly, bone marrow cells were harvested by cutting the femur of 8-10 week old mice at the knee joint and flushing with sterile ice-cold Phospate-buffered saline (PBS) using a 5-ml syringe and a

25-gauge needle. Cells were counted using hemacytometer and seeded at density of 2 ×

106 cells/ml in BMDM medium (complete DMEM medium supplemented with 10%

L929-conditioned medium.) in 10-cm petri dish. To prepare L929-conditioned medium,

1× 106 L929 cells were cultured in 10-cm plate containing 10 ml of DMEM medium in a humidified incubator with 5% CO2 at 37°C for 5 days. L929-conditioned medium was harvested and filtered through a 0.45-m filter (Millipore). Cells were maintained in a humidified incubator with 5% CO2 at 37° C for 4 days then washed with PBS and added with fresh BMDM medium. Differentiated cells were used for experiments at day 7. α-

Amanitin (Santa Cruz Biotechnology, Inc. CA) and Actinomycin-D (Sigma-Aldrich, MO) were used at a final concentration of 1 µg/ml.

2.2 Plasmids

The expression vectors for EPC MAVs and EPC IFN were obtained from Dr.

Michel Brémont (French National Institute for Agricultural Research [INRA], Jouy-en-

Josas Cedex, France). The pBK-ITS1 and L expression plasmids were from Dr. Vikram

N. Vakharia (University of Maryland, Baltimore County), the Tet-mSEAP construct was obtained from Dr. Fan Dong (University of Toledo) and the IFN-luciferase reporter obtained from Dr. John Hiscott (VGTI, Florida). The IFITM-1-luciferase, SV40- luciferase, CMV-lacZ, and CMV-GFP expression vectors have been described previously

(Pore, 2012). The expression vectors for human RNF114, RNF125, MDA5, RIG-I,

12 MAVs, TRAF3, TRADD, FADD, TBK1 were also previously cloned in our lab. IKK and NF-B luciferase construct were provided by Dr. Brian Ashburner (University of

Toledo). pLKO.1 puro vector (Addgene plasmid # 10879) was a gift from David Root, psPAX2 (Addgene plasmid # 12260) and pMD2.G (Addgene plasmid # 12259) were gifts from Didier Trono and purchased from AddGene. Plasmids expressing the VHSv viral proteins were previously cloned in our lab (Pore, 2012). VHSv M, NV, G, and N coding sequences were PCR amplified with appropriate primers with EcoR I and Kpn I site. All fragments were cloned into pcDNA 3.1 (-) myc/his A (Invitrogen), or p3xFLAG-CMV-

14 (Sigma). The mRNA seuquence used to design the luciferase primers was found on genbank (U47295). Primers and cloning sites for cloning were listed in table 1.

Luciferase driven by human U6 promoter reporter construct (pGL3-U6-Luc) was generated by subcloning human U6 promoter from pLKO.1 puro vector (Addgene plasmid # 10879) into pGL3 luciferase reporter vector (Promega #E1751) upstream of luciferase gene using Gibson Assembly (NEB #E5520). Rainbow trout pol I promoter luciferase construct (pGL3-ITS1-Luc) was generated by subcloning rainbow trout rRNA intergenic sequence region ITS1 into pGL3 luciferase reporter vector between two

HindIII sites. ITS1 promoter sequence and template were provided by Dr. Vikram N.

Vakharia (University of Maryland, Baltimore County).

2.3 VHSv Stocks and Infection

Stocks of the VHSv IVb strain initially were obtained from Dr. James Winton

(USGS, Seattle, WA.). VHSv F1strain was kindly provided by Dr. Gale Kurath (the

United States Geological Survey in Seattle, Washington). Recombinant WT VHSv GL,

VHSv DK-GL M and VHSv GL-DK M were generated from IVb strain MI03GL by Dr.

13 Vikram N. Vakharia (University of Maryland, Baltimore County). The virus was propagated by infecting a monolayer of EPC cells in serum free MEM media at

MOI=0.01 for 1 h and then culturing in complete MEM media at 20° C until complete cytopathicity was reached (typically 72 h). Media was collected and dead cells and debris cleared by centrifugation and filtration. Stocks were titered by serial dilution (1:3) on a

96 well plate of confluent EPC cells to determine viral concentration.

2.4 Cloning and PCR

The primers used to clone the VHSv, SVCV and SHRV M genes are listed in table 1. EPC cells were infected with VHSv for 48 h followed by RNA isolation. RNA was isolated using TRIzol per manufacturer’s protocol (Invitrogen, San Diego, CA). Two

µg of the RNA was reverse transcribed using M-MLV RT (Promega) as previously described (Sarela, 2000; Sambrook, Fritsch, & Maniatis, 1989). Briefly, two µg of RNA was mixed with 100 ng of random hexamer primers and the volume brought to 7 µl with water. Samples were incubated at 70°C for 10 min and the tubes briefly cooled before adding the M-MLV-RT mixture (4 µl M-MLV 5x Reaction Buffer, 2 l dNTPs

(Invitrogen), 0.5 l Recombinant RNasin Ribonuclease Inhibitor (Invitrogen), 0.5 l M-

MLV RT (Promega), and water to 25µl). The samples were mixed gently and then incubated at 42°C for 1 h. PCR was carried out with appropriate primer sets using annealing temperatures and elongation times found in Tables 1 and 2. PCR was started with a 5 min denaturation step at 95°C to activate the enzyme. Each cycle included a 30 sec 95°C denaturation step, 30-90 sec 54°C annealing step and 1 min per 1 kb 72°C elongation step.

14 2.5 Transfection

Transfections were performed by using Polyjet reagent (SignaGen, MD) following the manufacturer’s instructions. Briefly, plasmids were mixed with Polyjet in

SF medium for 20 min then added to cells. Media was then changed to complete medium after 3 h incubation.

2.6 Click-iT RNA Alexa Fluor 594 Imaging and Immunofluorescence Microscopy

Cells were seeded on poly-L lysine coated glass cover slips for 24 h at a density

~30%. Click-iT assays were performed using Click-iT RNA Imaging Kits (Invitrogen,

CA) according to the manufacturer’s instruction. Briefly, after treatment cells were incubated with 1 mM 5-ethynyl uridine (EU) for 2 h, fixed in 3.7% formaldehyde in PBS for 15 min at room temperature, rinsed with PBS, permeabilized in 0.5% Triton X-100 in

PBS for 15 min then rinsed once with PBS. Cells were incubated in Click-iT reaction cocktail for 30 min in the dark then washed with Click-iT reaction rinse buffer. For immunofluorescent staining, the cells were blocked for 30 min at room temperature (1%

BSA in PBS), then with primary antibody (in 1% BSA in PBS) for 1 hr. Cells were washed in PBS 3 times for 5 min each, then FITC conjugated secondary antibody (in

1%BSA in PBS) was added for 1 h at room temperature. After a PBS wash, the cover slips were mounted to slides with ProLong Gold Antifade Mountant with DAPI (Life

Technologies, CA) for 24 h, then imaged on an Olympus IX81 inverted fluorescent microscope.

2.7 Luciferase Assays/-gal Assays

15 Cells were transfected with the appropriate plasmids in a 12-well tissue culture plate at a density ~70% for 24 or 48 h. After media was removed, cells were washed with

PBS twice then lysed with 150 l Cell Culture Lysis Reagent 5X (diluted to 1X in water)

(Promega, WI) for 15 min on ice. Half of the lysate was used in to assess luciferase activity and mixed with 50 l luciferin (26.67 mM luciferin, 25 mM Glycyl-glycine, 1 mM DTT) and 50 l ATP solution (25 mM Glycyl-glycine, 4 mM ethylene glycol tetraacetic acid (EGTA), 15 mM MgSO4, 15 mM K2HPO4, 1 mM DTT, 2 mM ATP). The plate was read rapidly for luminescence using a SpectraMax plate reader. The other half of the lysate was used for beta-galactosidase activity determination using 50 l -gal buffer (60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 2 mM MgSO4, 2.6 M ortho-

Nitrophenyl--galactoside (ONPG), 3.2 l -mercaptoethanol). The mixture was incubated at 37°C until samples started to become yellow. The absorbance was then read at 414 nm on SpectraMax plate reader. In most cases, the luciferase reading was normalized to the -gal reading. To obtain a fold induction value, each sample was divided by the normalized value of the negative control. The exception was with M transfection studies, since M blocks SV40 beta-galactosidase expression (data not shown).

2.8 Immunoblotting

Cell lysates were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) as previously described (Lai et al., 2015). Briefly, samples were run on a 12.5% separating gel (2.5 ml 4x separating buffer, 3.125 ml 40% acrylamide, 100 l 10% APS, 10 l TEMED, 4.265 ml H2O) and 4% stacking gel (1.25 ml 4x stacking buffer, 0.625 ml 40% acrylamide, 50 l 10% APS, 5 l TEMED, 3.07 ml

16 H2O) in running buffer (25 mM Tris base, 192 mM glycin, and 6.94 mM SDS) at 100 volts until protein marker separated and reached the desired position. Proteins were transferred to Immobilon-P Polyvinylidene Fluoride (PVDF) membrane via semi-dry electrophoretic transfer and blocked in 1% BSA in TBST. Membranes were incubated with primary antibody dilutions (in 1% BSA in TBST) at 4°C overnight. The next day, membranes were washed in TBST then incubated with secondary antibody conjugated with HRP in 1% BSA in TBST for 1 hr at room temperature. Membranes were washed in

TBST and incubated with Enhanced Chemiluminescence reagent (Pierce) for 2 min then visualized on a UVP ChemiDoc-It2 510 Imager.

2.9 Real-time PCR

Quantitative real-time PCR was performed by mixing 10 l SsoFast Evagreen

Supermix master mix (Bio-Rad), 1 l cDNA, 50 ng of each primer and 8 l H2O (20 l reaction). PCR reactions were run on a Bio-Rad C1000 Thermal Cycler. The program used included denaturation at 95°C for 3 min followed by 40 cycles: 30 s at 95°C and 60 s at 60°C . Samples were normalized to human GAPDH, human -actin, mouse GAPDH, mouse -actin or fish -actin, depending on the design, and relative gene expression levels were calculated by using the ddCT method.

2.10 Lentivirus

RNF114 shRNA lentivirus was prepared following Addgene protocol. Plasmid pLKO.1 encoding control or RNF114 shRNA and packaging plasmids psPAX2 and pMD2.G were purchased from Addgene. HEK-293T cells were plated in 5 ml media

(DMEM + 10% FBS without antibiotics) in 60 mm tissue culture plate at density of 40%

17 overnight. Next day, the cells were transfected with plasmid cocktail (1 g pLKO.1 plasmid, 750 ng psPAX2, 250 ng pMD2.G, 6 l Polyjet then QS to 100 l with serum- free DMEM) for 15 h. Next morning, media was replaced with 5 ml complete DMEM medium and harvested after 24 h. This step was repeated once and total 10 ml media was harvested and spun down at 1,250 rpm for 5 min to remove cell debris. The supernatant containing lentivirus was stored at -80°C until use.

2.11 ChIP Assay

The chromatin immunoprecipitation (ChIP) assay was performed as previously described (Kallesen & Rosen, 2001) with the following modifications. Briefly, 107 cells were cross-linked with 1% formaldehyde for 10 minutes and then quenched with 125 mM glycine for 5 minutes at room temperature. Nuclei were prepared in Cell Lysis

Buffer (5 mM Tris-HCl pH 8, 85 mM KCl, 0.5% NP-40, 0.5 mM PMSF, 1X protease inhibitor cocktail (PIC, Thermo Scientific)) on ice for 10 minutes and sonicated to yield chromatin fragments (200 to 700 bp). Immunoprecipitations were performed overnight at

4 °C using 1 g of anti-pol II antibody A304-405A (Bethyl Laboratories, Inc., TX) or

IgG, and then incubated with protein A agarose (Millipore, Billerica, MA), which was pre-equilibrated with sonicated herring sperm DNA and BSA. Immunoprecipitated material was washed extensively, and the cross-links reversed. DNA from the eluted chromatin was purified by PCR purification kit following manufacture protocol (Qiagen).

Differences in DNA enrichment for ChIP samples were determined by qPCR using 4% of the precipitated sample DNA and 1% of the input DNA. The primers used for ChIP assay are listed in Table 1.

18 Table 1: Primers for PCR Analysis and Cloning

Restriction Primers Sequence Site mSEAP se GACCCTGCTCAGGACCCTC mSEAP as GATTTGCCATCCTCAGCCTTG U6 promoter se TCTCTATCGATAGGTACCTTTCCCATGATTCCT Kpn I TCATATTTG U6 promoter as CAGTACCGGAATGCCAAGCTTCGTCCTTTCCA Hind III CAAGATATATAAAG CMV se CGTTTAGTGAACCGTCAGATCG CMV as CCGGTGTCTTCTATGGAGGTCA IVb M se ACGAATTCATGGCTCTATTCAAAAGAAAGCGC EcoR I ACCATCCTG IVb M as ACGGTACCCCGGGGTCGGACAGAG Kpn I IVb M mid se ACAAGCTTCAAGATAGCTGAAGC Hind III IVb M mid as ACAAGCTTGTGATCAGGGTTTTG Hind III F1 se ACGAATTCATGGCTCTGTTCAAAAGAAAGCGC EcoR I ATCATCC F1 H as ACAAGCTTGGTACCCCGGGGCCG Hind III F1 K as ACGGTACCCCGGGGCCGGGCAGAGGGGG Kpn I D62A se TCTCTGTGAAGCTCAACATCCT D62A as AGGATGTTGAGCTTCACAGAGA SVCV M se CAGAATTCATGTCTACTCTAAGAAAG EcoR I SVCV M as CAGGTACCATCTCCCATGAACAGGGA Kpn I SHRV M se CAGAATTCATGGCAGAATCGATCGAG EcoR I SHRV M as CAGGTACCCTTTCTTGAGGACTCGTT Kpn I IHNV M se ACGAATTCATGTCTATTTTCAAGAGAGC EcoR I IHNV M as CTTGGTACCTTTTTCCTTCCCCCGCTTTTCGG Kpn I Virus down se ACGGATCCAAAACGCAGATCAG Virus down as AGGGGTGAGTATACAGTGGAGT VHS clone se AAGCTAGCACAAAAAACATGGCTCTATTCA Nhe I VHS clone as TTCAGCTGGTTGTGTACACAAA Pvu II hRNF114 se CCGAATTCCAAGATGGCGGCGCAACAG hRNF114 as GATGCGGCCGCTTCACTGGTCGATGATG mRNF114 se CATCCCAAACCGATACACCT mRNF114 as GTCGTAGGAAAAAGCGGTGACG hISG56 se TCACCAGATAGGGCTTTGCT hISG56 as CACCTCAAATGTGGGCTTTT mISG56 se CTCAGAGCAGGTCCAGTTCC mISG56 as TCCATCTCAGCACACTCCAG mIFNβ se TTCTCCAGCACTGGGTGGAA mIFNβ as AGGTACCTTTGCACCCTCCA mIL10 se ACCTGGTAGAAGTGATGCCCCAGGCA mIL10 as CTATGCAGTTGATGAAGATGTCAAA hActin se GTGCCCATTTATGAGGGCTA

19 hActin as CTGGCAGCTCGTAGCTCTTT hGAPDH se AAATCCCATCACCATCTTCC hGAPDH as GTCCACCACCCTGTTGCTGT mGAPDH se TTGTCAGCAATGCATCCTGC mGAPDH as TTGCCCACAGCCTTGGCAGC fActin se AGACATCAGGGTGTCATGGTTGGT fActin as GGGGTGCTCCTCTGGGGCAA

Chapter 3

VHSv Matrix Protein Inhibits Host Transcription

3.1 Introduction

3.1.1 Viral Hemorrhagic Septicemia

Viral hemorrhagic septicemia, historically known as Etgved disease (Jensen,

1965), is one of the most deadly infectious fish diseases, affecting more than 80 marine and freshwater fish species worldwide (Wolf, 1988; World Organisation for Animal

Health [OIE], 2015). Since first reported in freshwater-reared European rainbow trout

(Oncorhynchus mykiss) in the 1930’s by Schäperclaus (1938), VHS spread rapidly within

Europe over the next 20 years, causing several outbreaks with devastating consequences in rainbow trout farms (Pliszka, 1946; Scha¨perclaus, 1954; Rasmussen, 1965; Besse,

1955, Ross et al., 1994; Schlotfeldt et al., 1991). VHS was not detected outside of Europe until the late 1980s, when it was found in Chinook (Oncorhynchus tshawytscha) and coho salmon (O. kisutch) at two Washington State hatcheries in the US (Brunson et al., 1989;

Hopper 1989). Since then, VHS has been reported in Europe (Dixon et al., 1997;

21 Mortensen et al., 1999), Japan (Takano et al., 2000; Byon et al., 2006) and Korea (Kim et al., 2003).

The typical clinical signs of VHS include hemorrhages in the meninges, serous surfaces, skin, muscles, gills, fins, internal organs and eyes, exophthalmia, darkening of the body and pale gills. Other symptoms include ascites formation, abnormal behaviors or lethargy (Essbauer & Ahne, 2001). The disease is characterized by damage to the endothelial lining of blood vessels that primarily targets kidney and spleen (Yasutake

1975; De Kinkelin et al., 1979; Evensen et al., 1994). The VHS mortality rate can reach

100% in juvenile fish and 30-70% in older fish. Infected fish usually die from internal organ failure (Wolf 1988). Diseased fish can spread VHSv by shedding virus via urine and ovarian fluids. Surviving fish can become lifelong carriers that may not show clinical signs but still can release virus when stressed (Skall et al., 2005).

3.1.2 Viral Hemorrhagic Septicemia Virus

The causative agent of VHS is viral hemorrhagic septicemia virus (VHSv), which belongs to the Rhabdoviridae family and Novirhabdovirus genus (Walker, P., & Winton,

J. 2010). VHSv was first isolated by Jensen (1963) using trout cell cultures. VHSv is an enveloped, negative sense, single-stranded RNA (ssRNA) virus with a single, non- segmented RNA genome of approximately 11 kb (Fig. 3-1). The genome encodes six proteins in the following order (3’-5’): nucleoprotein (N; 44 kDa), phosphoprotein (P; 25 kDa), matrix protein (M; 20 kDa), glycoprotein (G; 63 kDa), nonvirion protein (NV; 13.7 kDa) and the viral polymerase protein (L; 224 kDa) (Schütze et al., 1999; Ammayappan

& Vakharia, 2009).

Nucleoprotein (N) Phosphoprotein (P)

Matrix protein (M)

Glycoprotein (G)

Polymerase (L)

N P M G Nv L

3’ 5’

Fig. 3-1. Schematic of VHSv Virion Structure and Genome. VHSv is a bullet shaped, envoloped rhabdovirus containing a negative sense, single stranded RNA genome that encodes 6 viral proteins. G is a transmembrane protein that forms homotrimer spike-like projections that extend out of the viral surface. The G protein is responsible for viral attachment and entry into host cells as well as interacting with the M protein. The N protein associates with viral RNA and forms ribonucleocapsid core of VHSv with M, P and L protein. The NV protein is not present in the viral particle.

23 The VHSv genome is packaged in a helical ribonucleocapsid core surrounded by a lipid bilayer envelope. The glycoprotein of VHSv forms homotrimer spike-like projections that extend outward through the lipid bilayer. The G protein is responsible for viral attachment and entry into host cells by interacting with cell receptors that mediate virus uptake by endocytosis. G protein can also induce host immune responses and is recognized by neutralizing and protective antibodies (Lorenzen N et al., 1999). The ribonucleocapsid core of VHSv consists of the RNA genome, N, P and L protein (Rose &

Whitt 2001). The function of the non-virion protein (NV) is not fully understood and NV is unique to VHSv and the other fish novirhabdovirus infectious hematopoietic necrosis virus (IHNV). Recent reverse genetics studies showed NV gene deletion in both IHNV and VHSv impaired viral growth in cell culture and had reduced pathogenicity while normal IHNV growth was restored by expressing NV in cells (Ammayappan et al., 2011;

Thoulouze, 2004). Replacing IHNV NV with VHSv NV also did not affect IHNV replication although sequence similarity is low between the two viral NV proteins

(Thoulouze, 2004). Another study implicated an antiapoptotic function of VHSv NV.

NV-deficient and NV knockout mutant VHSv virus induced earlier apoptosis than the wild-type recombinant virus and NV protein function was restored by replacement of

IHNV NV gene (Ammayappan & Vakharia, 2011). Nevertheless, the mechanism of Nv action remains unknown.

The life cycle of VHSv is similar to that of other rhabdoviruses. The glycoprotein initiates the absorption step by binding to cellular membrane receptors or extracellular matrix glycoprotein fibronectin (Bearzotti M et al., 1999). Several studies also suggest phosphatidylserine could be a binding target of G protein (Estepa et al., 2001; Estepa &

24 Coll. 1996; Nunez et al., 1998). Virus particles are internalized by endocytosis in coated vesicles (Granzow et al., 1997). The viral nucleocaspid core is released at low-pH (~6) via temperature-dependent (14°C ) membrane fusion between virus and primary lysosome mediated by G protein (Estepa & Coll 1997). After nucleocaspid release into the cytoplasm, positive sense mRNA of each VHSv gene is transcribed by the RNA dependent RNA polymerase complex comprised of L and P (Naito & Ishihama 1976).

The viral genes are separated by conserved gene junction sequences that are responsible for transcription stop-start, capping and polyadenylation (Barr et al., 1997; Hwang et al.,

1998). Viral genes are sequentially transcribed from 3’ to 5’ with one entry point at the 3’ end of the genome (Ball & White 1976; Abraham & Banerjee 1976). Due to transcriptional attenuation, the polymerase is not able to reinitiate transcription and sometimes disassociates from the genome at gene junctions (Iverson & Rose 1981). This results in more mRNA product of genes that are closer to the 3’ end, such that the relative expression levels of each gene is regulated. When sufficient N protein expression is attained to cover the nascent RNA, L protein switches its function from transcription to replication (Arnheiter et al.,1985; Vidal & Kolakofsky,1989). The full length, positive strand antigenome is transcribed and serves as template to synthesis new viral genome

(Leppert et al., 1979). G protein is processed in endoplasmic reticulum (ER) and golgi apparatus then transported to plasma membrane. M proteins associate with the lipid bilayer and the inner portion of the G protein functions as a link between G and viral nucleocaspid (Dancho et al., 2009; Mebatsion et al., 1999) to assemble virions and initiate the budding process (Jayakar et al., 2000).

3.1.3 Nomenclature and Classification

25 VHSv isolates are grouped into 4 strains (designated I to IV) based on phylogenetic analysis of the G and N genes (Einer-Jensen et al., 2004; Snow et al., 2004).

Each group affects specific geographic regions and has unique host specificities. Strain I is primarily found in Europe and is further divided into five substrains (designated Ia-Ie)

(Thiery et al., 2002; Einer-Jensen et al., 2004). Ia is responsible for most outbreaks in

European freshwater rainbow trout farms (Einer-Jensen et al., 2004; Snow et al., 2004;

Stone et al., 2008; Toplak et al., 2010), but was also reported in brown trout (Thiery et al.,

2002), pike (Meier & Jørgensen 1979), grayling and one marine isolate from German turbot (Schlotfeldt et al., 1991; Wizigmann et al., 1980). The Ia substrain is further separated into two clades: the Danish Ia-1 and Ia-2 from other European countries (Kahns et al., 2012). The Ib substrain arose from marine isolates originating from the Baltic Sea,

Skagerrak, Kattegat, North Sea and the English Channel (Dixon et al., 1997; Mortensen et al., 1999; Batts & Winton 2007; Takano et al., 2000). Substrain Ic viruses were isolated from farmed rainbow trout in Denmark (Nishizawa et al., 2002), while Id comprises a group of isolates from Norway, Finland and the Gulf of Bothnia sea-reared rainbow trout (Snow et al., 1999; Raja-Halli et al., 2006). Genotype Ie isolates are found in Georgia and Turkey (Snow et al., 1999; Ogut & Altuntas 2014). Strain II viruses are isolated from the Baltic Sea region (Mortensen et al., 1999). Strain III are reported from the North Sea (Smail 2000), Ireland (Snow et al., 1999), Skagerrak (Mortensen et al.,

1999), France (Jørgensen et al., 1994), Norway (Dale et al., 2009), Flemish Cap (Dopazo et al., 2002) and Japan (Isshik et al., 2001). Strain IV is divided into three substrains. IVa was first identified as including marine isolates from Japan, Korea and the western coast of the United States (Kim et al., 2003; Nishizawa et al., 2002). In 2005, a novel substrain

26 of VHSv was isolated in muskellunge (Esox masquinongy) from Lake Ontario and was also found in an archived sample from Lake St. Clair, dating back to 2003 (Elsayed et al.,

2006). Genotyping results suggested this new virus (MI03GL) was closely related to

Strain IV yet differed from previous isolates. Therefore a new sublineage IVb was established. Recently a new substrain was identified, designated as IVc (Pierce & Stepien

2012) including isolates from mummichog (Fundulus heteroclitus), brown trout (Salmo trutta), stickleback (Gasterosteus aculeatus aculeatus) and striped bass (Morone saxatilis;

Gagné et al. 2007).

3.1.4 VHSv IVb in the Great Lakes

Since the first VHS endemic was reported in the Great Lakes region, VHSv IVb has been isolated from 31 fish species including muskellunge, freshwater drum, lake trout, chinook salmon, yellow perch, walleye and others (Thompson et al., 2011). Several epizootics occurred during the next decade causing massive die-offs among many freshwater species and posing a threat to the fish farming and sport fishing industries

(Lumsden et al., 2007; Groocock et al., 2007; Gagné et al. 2007). By 2010 VHSv has been detected in all five of the Laurentian Great Lakes (Thompson et al., 2011; Cornwell et al., 2011). The IVb substrain is closely related to the marine IV strain and presumably evolving from marine strains by adapting to a freshwater environment (Pierce & Stepien

2012). Despite intensive management and surveillance, VHSv is till detectable among asymptomatic fish in the Great Lakes region (Kim & Faisal 2011).

3.1.5 Transmission

27 VHSv spreads from fish to fish primarily through lateral transmission. Surviving fish can become lifelong carriers as viral reservoirs shedding virus in urine and reproductive fluids (Smail & Snow 2011). Host gill epithelium, fin base or skin are likely the main entry point for VHSv (Chilmonczyk et al., 1995; Yamamoto et al., 1992;

Harmache et al., 2006). Oral transmission by ingestion of infected prey has also been reported (Ahne 1980; Schönherz et al., 2012). The optimal temperature for VHSv is between 9-12°C (Goodwin & Merry 2011) and temperature above 20°C reduces viral pathogenicity (De Kinkelin 1970).

Although much has been learned about VHSv transmission and spread, little is known about the details of viral/host interaction, including the determinants of viral detection and/or host suppression. The following studies were initiated to bridge this gap and start to uncover cellular pathways impacted during VHSv infection.

3.2 Results:

3.2.1 VHSv Inhibits Host Antiviral Responses

3.2.1.1 VHSv IVb M Inhibits Host Gene Expression

A previous study in our laboratory showed that upon infection, VHSv produces a factor or factors that are capable of shutting down host IFN-mediated antiviral responses, and that viral infection potently suppressed IFN responsiveness (data no shown. Pore,

2012). These data supported the idea that VHSv is capable of inhibiting host cell production and/or response to IFN.

To identify viral proteins involved in the observed inhibitory effect of VHSv on host IFN responsiveness, each of the six VHSv structural and nonstructural genes was cloned into an eukaryotic expression plasmid and then co-transfected along with

IFITM1/luc and fish MAVS (fMAVS). Ectopic fMAVS overexpression is sufficient to induce endogenous IFN expression, which feeds back on the cells to activate the IFITM1 promoter and induce luciferase expression. Although several of the viral genes had subtle effects on IFITM1 induction, only the pcDNA3.1 M plasmid (pCD-M) potently decreased IFITM1 promoter activation (Fig. 3-2A). To determine whether the impact of

M was upstream or downstream of IFN expression, cells were transiently transfected with pCD-M and IFITM1/luc and then treated with fIFN (Fig. 3-2B). M once again dramatically inhibited IFN-induced IFITM1 promoter activity. For anlaysis of pathways upstream of IFN, EPC cells were co-transfected with fMAVS, pCD-M and an IFN

29 promoter-luciferase reporter plasmid (IFN/luc). M also potently blocked induction of luciferase activity in a dose-dependent manner (Fig. 3-2C), with 80% inhibition observed at only 0.5 ng of pCD-M/5X105 cells. The equally potent inhibition of both IFN and ISG transcriptional induction suggested either that M exhibited multiple effects and targeted components of both pathways, or that it was a general inhibitor of gene transcription or translation. To test this possibility, pCD-M was co-transfected with an unrelated, constitutively active SV40 promoter-luciferase reporter (SV40/luc). VHSv M inhibited

SV40 promoter activity in a dose-dependent manner (Fig. 3-2D) suggesting that M inhibits a general step in cellular protein expression – either transcription or translation.

In order to clarify whether M functioned as a transcriptional or translational inhibitor, cells were again co-transfected with pCD-M and SV40/luc plasmids and luciferase mRNA quantified by using real time RT PCR. M co-expression induced a dose- dependent decrease in luciferase mRNA expression, strongly implicating an impact on cellular RNA transcription or half-life. Because the impact of M on luciferase mRNA was not as potent as that observed when measuring luciferase activity (Fig. 3-2A compared to 3-2E) we reasoned that M either exhibited secondary effects on mRNA translation, or that luciferase mRNA expression begins in transfected cells before M translation can elicit an inhibitory effect (Fig. 3-2E). Thus, in order to measure the impact of M on transcriptional initiation instead of pre-existing steady mRNA levels, we cotransfected pCD-M with a plasmid encoding a mouse SEAP (secreted embryonic alkaline phosphatase) reporter gene under the transcriptional control of a tetracycline responsive element. After 24 h, transfected cells were left unstimulated or were treated with doxycycline for 24 h prior to RNA extraction and real time RT PCR analysis of

30 SEAP mRNA levels. Under these conditions, SEAP mRNA induction was completely inhibited by M co-expression (Fig. 3-2F). Taken together, these data suggest that VHSv

M inhibits host cellular transcription during viral infection.

31 A

t i 200

t c

e 150

c 100

i 50 l

o 0

IFITM1/Luc + + + + + + + + + + + + + + fMAVS - + + + + + + + + + + + + + .05 .1 .05 .1 .05 .1 .05 .1 .05 .1 .05 .1 M P Nv N G L

ng plasmid/5x105 cells

t i 150

a 100

d 50

r o 0

N IFITM1/Luc + + + + + + fIFN - + + + + + IVb (MI03GL) M - - .5 1 10 50 ng plasmid/5x105 cells

32 C

t i 800

e 600

c 400 u

i 200

l a

m r o 0 N hIFN/Luc + + + + + + fMAVS + + + + + + IVb (MI03GL) M - .5 1 5 10 50 5 ng plasmid/5x10 cells

t i 150

e s a 100

i c

d 50

r o 0

N SV40/Luc + + + + + + IVb (MI03GL) M - .5 1 5 10 50 5 ng plasmid/5x10 cells

33 E

v 150

R 100

e s

c 50

a l 0 e

R SV40/Luc - + + + + - IVb (MI03GL) M - - .1 .2 .5 .5 6 g plasmid/10 cells

s l 60 e ****

N 40

E S 20

R 0

Tet-mSEAP + + + Dox - + + IVb (MI03GL) M - - +

34 Fig. 3-2. VHSv-IVb M Inhibits Host Gene Expression. A) EPC cells were transfected with 0.4 µg IFITM1/luc construct. Fish MAVS (0.3 µg) and plasmid encoding various virus genes (0.05 or 0.1 µg) were co-transfected as indicated followed by luciferase assay

48 h later, where luciferase activity of IFITM1/luc + MAVS is set at 100. B) IFITM1/luc

(0.4 µg ) and VHSv IVb M (0.5-50 ng) were co-transfected in EPC cells for 24 h then treated with or without fIFN for 24 h followed by luciferase assay. Luciferase activity of

IFITM1/luc + IFN is set at 100. C) IFN/luc construct (0.4 µg), fMAVS (0.3 µg) and

VHSv IVb M (0.5-50 ng) were co-transfected into EPC cells for 24 h followed by luciferase assay. Luciferase activity of IFN/luc + MAVS is set at 100. D) SV40/luc construct (0.4 µg) and VHSv IVb M were co-transfected into EPC cells for 24 h followed by luciferase assay. Luciferase activity of SV40/luc is set at 100. E) SV40/luc construct

(0.8 µg) and VHSv IVb M were co-transfected into EPC cells for 24 h. Luciferase mRNA levels were measured by qRT-PCR. Data were normalized to fish actin mRNA levels.

(experiment was performed by Dr. Douglas Leaman.) F) Tet-mSEAP were co-transfected with or without IVb M (0.1 µg) into EPC cells for 24 h then treated with doxycycline for

24 h followed by RNA isolation and qRT-PCR using mSEAP primer. **** p<0.001.

35 3.2.2 M Protein Inhibits Host Transcription

3.2.2.1 VHSv M Blocks Nascent Cellular RNA Synthesis

Previous studies on the anti-host role of vesicular stomatitis virus (VSV) M protein demonstrated direct inhibition of host transcription (Yuan et al., 1998). To determine if VHSv M behaved similarly, we utilized an analog of uracil, 5-ethynyl uridine (EU), to study active (nascent) cellular RNA synthesis. EU contains an alkyne that can react with an azide-modified fluorophore to give fluorescent signal (Jao & Salic,

2008; Salic & Mitchison, 2008). EPC cells were left untreated, treated with α-Amanitin

(1 ug/ml), actinomycin D (1 ug/ml) or infected with VHSv IVb (moi = 1) for 24 h then pulsed with EU for 2 h, labeled with the Click-iT reagent and imaged. Untreated, labeled cells showed nuclear RNA staining with strong puncta representing rRNA synthesis. α-

Amanitin, which inhibits predominantly RNA polymerase (RNAP) II-mediated transcription at the dose used (Lindell et al., 1970), showed strongly reduced nuclear labeling but with little inhibition of rRNA nucleolar staining (Fig. 3-3A). In contrast,

Actinomycin D, which inhibits RNAP I, II and III, and thus served as a control for complete transcriptional inhibition, suppressed all RNA synthesis in treated cells (Fig. 3-

3A). EU staining in VHSv infected cells mimicked the pattern observed with α-Amanitin treatment, but residual nucleolar staining in VHSv infected cells was demonstrably less than in α-Amanitin treated cells (Fig. 3-3A). These data suggested that VHSv potently inhibited activity of all three RNA polymerases to varying degrees. To assess a direct role for M in the observed virus-dependent inhibition of host transcription, we cotransfected pCD- M with a CMV-GFP plasmid (as a marker of transfection) into EPC cells for 24 h and then labeled cells with 5-EU. M transfected cells exhibited decreased nascent cellular

36 RNA staining as compared to GFP-only transfected control cells (Fig. 3-3B). The reduction mirrored that observed with viral infection, implicating M as a contributing viral protein.

37 A DAPI VHSV Nacent RNA

Negative

Positive

- Amanitin

ActinomycinD

IVbVHSv

38 B Control IVb M

Nascent RNA

GFP

DAPI

39 Fig. 3-3. VHSv M Blocks Nascent Cellular RNA Synthesis. A) EPC cells were left untreated, treated with 2 µg/ml α-Amanitin or infected with 1 m.o.i VHSv IVb for 24 h then fed with EU for 2 h. EU was stained with Alexa594 via click chemistry reaction.

VHSv is probed with polyclonal anti-VHSv antibody. B) EPC cells were cotransfected with 0.4 µg GFP and 0.1 µg IVb M for 24 h then labeled with 100 µM EU for 2 h. EU was stained with Alexa594 via click chemistry reaction.

40 3.2.2.2 VHSv IVb M Inhibits RNAP I-III

To more precisely define which transcriptional responses might be inhibited by M, we tested VHSv M inhibitory potency on RNAP I, II and III-regulated promoters in a cell-based luciferase assay. The SV40/luc reporter used previously was used again as an indicator of pol II-dependent transcription. A human U6/luc reporter (subcloned from pLKO.1) was used as a surrogate for to monitor pol III-dependent transcription, and a rainbow trout rRNA intergenic sequence region (ITS-1/luc) reporter was used to assess pol I-dependent transcription. M inhibited all three promoters’ activities, but pol II promoter appears to be more sensitive to the inhibitory effects of M than pol I and pol III promoters (Fig. 3-4A). Previous studies had implicated VSV M in suppression of all three RNA polymerases with differing efficacies (Ahmed & Lyles. 1998), and our studies suggest that VHSv IVb M is similarly effective in blocking pol I-III dependent transcription.

In addition to direct inhibition of cellular transcription, previous studies on VSV

M protein had implicated a role in blocking nuclear export of mRNAs (von Kobbe et al.,

2000). To determine whether VHSv infection alters the subcellular distribution of cellular mRNAs, EPC cells (5X106) were left uninfected or were infected with VHSv IVb strain

(moi = 1) for 24 h. Cells were separately treated with Leptomycin B for 3 h as a control for nuclear export inhibition. After treatment or infection, cells pellets were spiked with

1X105 human (HEK 293) cells to serve as an internal control. The cell mixtures were then separated into nuclear and cytoplasmic fractions and RNA isolated from each. Real time RT PCR was then performed to assess the subcellular distribution of fish actin mRNA in the uninfected, infected or leptomycin B treated cells. To control for variability

41 in fractionation fidelity or RNA extraction efficiency, human GAPDH mRNA was quantified in parallel and used to normalize the fish actin mRNA values. Since the human cells had not been infected or treated with leptomycin B, the human GAPDH mRNA distribution in the spiked 293 cells was expected to be identical in each sample, and the human primers were designed and validated not to cross react with fish GAPDH cDNA

(data not shown). Our data showed that VHSv infection resulted in a decrease in overall actin mRNA levels, but had only a moderate impact on the nuclear to cytoplasmic ratio as compared to uninfected cells (Fig. 3-4B). In contrast, leptomycin B altered dramatically the proportion of mRNA in the nucleus relative to the cytoplasm (Fig. 3-4B). From these data we cannot rule out a minor role for M in altering mRNA subcellular localization, but its primary anti-host effect appears to be the inhibition of nascent transcription.

42 A

SV40 (pol II)/Luc ITS1 (pol I)/Luc U6 (pol III)/Luc

140

120

100

control to relative percentage 20

0 IVb M - + - + - + 5 10 ng plasmid/5x10 cells

43 B

ns 1.5

L ****

N 1.0

0.5

e R 0.0 CTL MI03GL LMB

r 2.0

m *** s

p **

o t 1.5 y

u 1.0

n 0.5

m 0.0

c CTL MI03GL LMB

44 Fig. 3-4. VHSv-IVb M Blocks pol I-III Dependent Transcription. A) EPC cells were transfected with SV40-, ITS1- or U6-Luc reporter construct with M for 24 h followed by luciferase assay. B) EPC cells were left uninfected or were infected with VHSv IVb virus

(moi=1) for 24 h or treated with Leptomycin B (LMB) for 3 h serviced as positive control for nuclear export inhibition. Cells were spiked with 1X10^5 HEK 293 cells before fractionation followed by RNA isolation and qRT-PCR. Total fish actin mRNA levels and nucleus to cytoplasm ratio of fish actin mRNA were calculated. Realtime PCR results were normalized to human GAPDH values. * p<0.05; ** p<0.01; *** p<0.005; **** p<0.001.

45 3.2.2.3 M Disrupts RNAP II Activity and Recruitment

To gain further insight into the mechanism by which M inhibits host transcription, we assessed the impact of M on recruitment of RNAP II to a core promoter by performing ChIP analysis. When TRE-mSEAP was cotransfected with M and then induced by doxycycline 24 h later, we found that RNAP II recruitment in both uninduced and induced cells was significantly decreased in the presence of M (Fig. 3-5A). These data suggest inhibition of transcription occurred at the most basic level, the recruitment of pol II to the core promoter.

The C-terminal domain (CTD) of RNAP II largest subunit (Rpb1) consists of 25 to 52 tandem copies of a conserved heptapeptide repeat with the consensus YSPTSPS

(Corden, 1990). During elongation, the CTD of RNAP II is progressively phosphorylated by different kinases (O'Brien et al., 1994). The CTD is phosphorylated predominantly at

Ser 2 and Ser 5 residues, and these modifications indicate different phases of transcription and RNAP II activity (Bensaude et al., 1999; Komarnitsky et al., 2000).

Early in the transition from preinitiation to elongation, the CTD is primarily phosphorylated on Ser5 residues. During elongation, phosphorylation occurs mainly on

Ser2 residues to generate elongation-proficient RNAP II. Finally, near the 3’ end of the gene, CTD phosphorylation is predominantly on Ser2 residues. To address whether

VHSv impacts RNAP II phosphorylation patterns, we infected EPC cells with VHSv for up to 72 hr. Given the observed decrease of RNAP II recruitment caused by M protein, we hypothesized that less RNAP II would enter the elongation phase. As expected, the phosphorylation level at Ser2 residue within the CTD of RNAP II was decreased during

VHSv infection (Fig. 3-5B). These data suggest that VHSv IVb M protein inhibits host

46 cellular transcription to suppress host immune responses by disrupting RNAP II activity and/or recruitment.

47 A

T 15

e 10

e 5 *** g

o 0 F Tet-mSEAP + + + + IVb (MI03GL) M - + - + Dox - - + +

TRE CMV mSEAP

Ctrl 12 24 36 48 72 h

S2 RNAP II

RNAP II

actin

VHSv-M

48 Fig. 3-5. M Disrupts RNAP II Activity and Recruitment. A) EPC cells were transfected with a tet regulated SEAP plasmid and pCD-M for 24 h and were then left untreated or were induced with 2 µg/ml doxycycline for 4 h. After crosslinking, chromatin was immunoprecipitated with antibodies to IgG and RNAP II and analyzed by qPCR using primers specific for the minimal CMV promoter. RNAP II recruitment was normalized to IgG control. * p<0.05; ** p<0.01; *** p<0.005. B) EPC cells were infected with VHSv IVb virus (moi=5) for 0-72 h. Cell lysates were separated by PAGE and immunoblots probed with antibodies recognizing RNA polymerase II CTD repeat

YSPTSPS (phospho Ser2), total RNAP II, actin and VHSv, followed by HRP secondary antibodies and chemiluminescent detection.

49 3.2.3 Single Amino-Acid Change in M Affects Its Inhibitory Function

M proteins from various VHSv strains and substrains, as well as from the related fish rhabdoviruses IHNV, SVCV and SHRV were tested for antitranscriptional activities in cell-based luciferase inhibition assays. Each M gene was cloned into pcDNA3.1 and co-transfected with SV40/luc into EPC cells for 24 h, after which time luciferase assays were performed. Like VHSv IVb M, all rhabdoviral M proteins inhibited luciferase expression to varying degrees, although SVCV and SHRV M exhibited less potent activities as compared to the VHSv and IHNV M proteins (Fig. 3-6A). Although most

VHSv substrain M proteins had similar activities to IVb M, one exception was a variant

M cloned from VHSv Ia substrain (F1). This M protein exhibited significantly less inhibition as compared to the IVb substrain M (Fig. 3-4A). This M variant differed from

IVb M at only four amino acid positions (T9I, D62G, E181A and V198A). In order to determine which of these changes impacted anti-transcriptional activity, we mutated these same residues in various combinations within the VHSv IVb M background. The two residues that impacted activity the most were at 62 (DG) and 181 (EA). Reverse mutations (G62D, G62D and A181E) within the Ia (F1) backbone resulted in antitranscriptional efficacy that was similar to IVb M (Fig. 3-6B), while IVb M protein mutated singly at position 62 (D62A) or doubly at 62 and 181 (D62A and E181A) each exhibited about 90% reduction in efficacy (Fig. 3-6C). These data implicate D62A as the dominant change, but also suggest that the 181 mutation may augment the impact of the

D62 amino acid substitution. Although overall sequence identity between VHSv and

VSV M proteins is low (Fig. 3-6D), the predicted secondary structure of both M proteins is conserved (Fig. 3-6D). The aspartic acid at position 62 is also highly conserved among

50 most VSV and VHSv strains (Gaudier et al., 2002). Based on the crystal structure of VSV

M (Gaudier et al., 2002), and the predicted structure of VHSv M (Fig. 3-6E), this aspartic acid is located in the loop between α1 and α2 on the surface of the molecule, and is spatially close to E181. Taken together, these data suggest that this aspartic acid residue may be crucial for M protein folding or interaction with itself or other target molecules.

51 A

700 600 500

400

RLU 300 200 100

SV40/Luc - + + + + + + + + +

KRRV SHRV

Bog

IVb SVCV IHNV

150 9

D a 100

T G

I 50 I

N 0 SV40/Luc + + + + + + + + + + + + M - 1 10 1 10 1 10 1 10 1 10 10 ng plasmid/5x105 cells

52 C

t 150 ** i

a 100 r *** e

d 50

r o N 0

SV40/Luc + + + + + + + IVb M - 1 10 - - - - D62A - - - 1 10 - - D62A E181A - - - - - 1 10 ng plasmid/5x105 cells

53 Fig. 3-6. Single Amino-Acid Change in M Affects Its Inhibitory Function. A) M proteins from various VHSv strains and substrains (IVb: MI03GL; F1: Egtved isolate of

Ia strain from Denmark; Bog: NA-5 isolate from Bogachiel River; EB: EB#7 isolate from

Elliot Bay; Bog KRRV: isolate of IVa strain from Japan), and from IHNV, SVCV and

SHRV were co-transfected with SV40-Luc into EPC cells for 24 h, after which time luciferase assays were performed (experiment was performed by Tyler Williams). B)

EPC cells were co-transfected with various M mutants with SV40-Luc for 24 h followed by luciferase assay. C) EPC cells were co-transfected with M mutants (D61A, D62A and

E181A) with SV40-Luc for 24 h followed by luciferase assay. * p<0.05; ** p<0.01; *** p<0.005; **** p<0.001. D) VHSv and VSV M sequence alignment and predicted secondary structure using Clustal Omega program. α-helix (red) and β sheet (green) were shown in image. E) VHSv M predicted crystal structure with D62 location using Swiss

PDB Viewer 4.1.0 performed by Wade Weaver.

54 3.3 Discussion

Most if not all viruses have evolved mechanisms to inhibit the expression or function of host innate immune genes during virus replication (Holland & Peterson, 1964;

Wertz & Youngner, 1970). Inhibition of host transcription is a common strategy used by

RNA viruses that replicate entirely within cytoplasm, as a means of enhancing viral resources and reducing immune responses. Shutting down host transcription not only frees up cellular translational machinery that can be used for biosynthesis of viral gene products, it also inhibits the host antiviral responses by preventing synthesis of antiviral proteins. One clear example of an anti-host protein found within the Rhabdovirus family is the VSV M protein. VSV M potently inhibits host gene expression, thereby suppressing host antiviral responses, including type I IFNs upregulation (Ferran & Lucas-

Lenard, 1997; Ahmed et al., 2003). The VSV M protein also may block mRNA export from the nucleus (Clinton et al., 1978; Petersen et al., 2000; von Kobbe et al., 2000).

These anti-host functions of VSV M are separate from its functional role in viral assembly and budding (Dancho et al., 2009; Mebatsion et al., 1999; Black et al., 1993).

The fish rhabdovirus infectious hematopoietic necrosis virus (IHNV) M protein also inhibits host-directed gene expression (Chiou et al., 2000).

Here, we report that VHSv M protein similarly blocks host cellular transcription, leading to the inhibition of host antiviral responses. We show that VHSv infection suppressed host IFN-mediated antiviral responses, and that expression of matrix protein alone potently inhibited both MAVS-mediated IFN expression and IFN responsiveness in cell-based luciferase assay. M also inhibited a constitutively active SV40 promoter function, suggesting that VHSv M acts similarly to other rhabdoviral matrix proteins by

55 shutting down general transcription or translation. This possibility was enforced and clarified by the observation that VHSv infection or ectopic expression of M led to decreased host nascent RNA transcription. Our results support the hypothesis that VHSv

M protein inhibits host transcription as a means of promoting viral dissemination.

Previous work showed that VSV M protein was capable of blocking host transcription directed by all three host RNA polymerases (Ahmed & Lyles, 1998). We tested the inhibitory potency of VHSv M on three different promoters using luciferase constructs driven by the pol I-dependent Atlantic salmon ITS1 promoter, the pol II- dependent SV40 promoter and the human pol III-dependent U6 promoter, respectively.

Our data were consistent with previous observation made with VSV M, in that VHSv M inhibited transcription mediated by all there RNA polymerases, albeit to varying degrees.

Interestingly, EU staining of nascent RNA synthesis in VHSv-infected and M-transfected cells resembled the pattern of α-Amanitin treatment. Since α-Amanitin targets pol II and

III, but not pol I, this suggested that perhaps M targets all three host RNAPs through a common mechanism, but that this mechanism is not equally efficacious against all three polymerases. One could certainly envision that continued rRNA syntheses would benefit the virus, so it is possible that residual pol I activity is intentional.

Previous studies of VSV M suggested that the TATA-binding protein (TBP) subunit of TFIID was a potential target of M, as TFIID isolated from VSV infected cells was inactivated in an in vitro transcription assay while the transcription activity could be reconstituted by adding purified recombinant TBP (Yuan et al., 1998). Our ChIP assay results implicate an M-dependent suppression of both basal and doxycycline-induced recruitment of RNAP II to the minimal CMV promoter region of a Tet-mSEAP reporter

56 gene. Taken together, our data suggest that VHSv M inhibited RNAP II-directed transcription at initiation step by interrupting RNAP II promoter binding presumably by targeting one or more basal transcription factors, a hypothesis currently being tested with targeted co- immunoprecipitation studies.

M proteins from a variety of VHSv strains and some closely related fish viruses, including IHNV, snakehead rhabdovirus and spring viremia of carp virus were all able to inhibit SV40 promoter activity, an observation consistent with the argument that suppressing host protein expression is a conserved role of M protein among different rhabdoviruses. Interestingly, an M clone isolated from a VHSv F1 strain (Ia substrain) sample was less potent than other M proteins in inhibiting transcription (Fig. 3-7A).

When comparing this F1 M sequence with that of WT IVb M, only four amino acids changes (T9I, D62G, E181A and V198A) were present. When we tested a range of targeted mutations at these positions, we found that G62D conversion in the Ia M clone enhanced function to approximately that of the IVb M (Fig. 3-7B). To further investigate the impact of the M alternatives, two mutant M constructs (D62A and D62A+E181A) were made within the IVb M background. Both exhibited decreased ability to inhibit cellular gene expression as compared to WT IVb M (Fig. 3-7C). The predicted secondary structure of VHSv M is quite similar to that of VSV M even though amino acid conservation between the two is low (Fig. 3-7D). The aspartic acid residue at position 62 is conserved across multiple VSV and VHSv strains. In the VSV M protein, this aspartic acid is located between helices α1 and α2, and is exposed on the surface of the protein

(Gaudier et al., 2002). These results indicate that this conserved aspartic acid residue may play a critical role in the anti-host function of M by serving a structural role of as a

57 binding site for protein-protein interactions with host factors involved in regulation transcription, or as a critical structural determinant that is required for proper folding.

Additional work is needed to separate these related possibilities.

In summary, our studies verify that, like most viruses, VHSv suppresses host innate immune responses. As with VSV, VHSv actively blocks host transcription to suppress host antiviral responses, including IFN and ISG expression, by inhibiting cellular transcription through the M protein. Our results suggest that VHSv M interrupted association of RNAP II with a basal gene promoter, leading to decreased RNAP II recruitment and mRNA elongation. VHSv M inhibited all three cellular RNA polymerases, suggesting functional consistency with VSV M. However, further investigation is required to determine whether host general transcription factors, i.e.,

TFIID or TBP, are the targets of VHSv M. We identified a naturally occurring M variant within the VHSv Ia substrain, D62G which dramatically decreased M anti-host efficacy.

Reverse genetics studies are in progress to introduce these mutations into the VHSv IVb backbone to see how they impact virulence within the context of this particular virus.

These studies should enhance our understanding of M’s role in host inhibition and, ultimately, viral replication. If we could identify an M mutant that lost all or part of its anti-host effects without losing viral replication potential, such mutants may show promise as candidates for construction of attenuated viral vaccines. This is ultimately a long-term goal of this work.

Chapter 4

RNF114 inhibits RLH-dependent antiviral responses

4.1 Introduction

4.1.1 Regulation of RLH pathway by Ubiquitination

Although activation of the innate immune system is necessary to prevent the spread of infection and to activate the adaptive immune response, tight regulation of

IFN and other proinflammatory cytokine production is also essential to prevent damage to the host caused by long-term or enhanced immune responses. Ubiquitination and deubiquitination are common post-translational modifications that play important roles in

RLHs signaling pathway regulation.

Ubiquitin (Ub) is a 76-amino acid (~8.5-kDa) polypeptide ubiquitously expressed across different tissues and organisms (Ciechanover 2005).

Ubiquitination is a common post-translational modification that covalently attaches an ubiquitin molecule to a substrate through a three enzymes system (Pickart 2001). First,

59 ubiquitin is attached to ubiquitin-activating enzyme (E1) by a thioester linkage via ATP- dependent activation. Next ubiquitin is transferred to an ubiquitin-conjugating enzyme

(E2) via transthioesterification. E3 ubiquitin ligases (E3) interact with Ub-loaded E2 and specific substrate to transfer ubiquitin to a lysine or N-terminus of the substrate or another ubiquitin molecule. This process can be reversed by deubiquitinating enzymes

(DUBs; Hurley et al., 2006). So far only two E1 have been identified in humans

UBA1/UBE1 and UBA6/UBE1L2 (Pelzer et al., 2007) while more than 50 E2s and 700

E3s exist. Ubiquitin can form polyubiquitin chains through one of seven lysine residues:

K6, K11, K27, K29, K33, K48 and K63, or it can form linear polyubiquitin chains through N-terminus and C-terminus intermolecular association (Kirisako et al., 2006).

Different polyubiquitin structures can serve distinct functions. K48-linked or K11-linked polyubiquitination is involved primarily in cellular protein proteasomal degradation

(Chau et al., 1989; Jin et al., 2008). K63-linked polyubiquitination usually plays a role in protein kinase activation, membrane trafficking, DNA repair and stress response

(Arnason & Ellison, 1994; Chen & Sun, 2009; Hofmann & Pickart, 2001; Johnson, 2002;

Spence et al., 1995).

Ubiquitination plays multiple essential roles in the RLH signaling pathway. The

TRIM25 E3 ubiquitin ligase, an innate immune gene, mediates the formation of K63- linked polyubiquitin chains on aa K172 of RIG-I, thereby stabilizing the interactions between RIG-I and MAVS to increase downstream signaling and augment antiviral responses (Gack et al., 2007, 2008; Shigemoto et al., 2009; Zeng et al., 2010). RING

(really interesting gene) finger protein RNF135, or Riplet, also targets RIG-I for K63- linked polyubiquitination within both the CARD and RD domains to activate RIG-I

60 signaling pathway (Gao et al., 2009; Oshiumi et al., 2009, 2010). The ubiquitin-editing protein A20 negatively impacts RIG-I-mediated IRF3 and NF-B signaling (Lin et al.,

2006), presumably via degradation of K63-polyubiquitin chains. TRAF3 promotes

MAVS-mediated RLH pathway activation by conjugating K63-linked polyubiquitin chains to MAVS, allowing recruitment of the downstream molecule IKKHäcker et al.,

2006; Oganesyan et al., 2006. RNF125, also named TRAC-1, an E3 ubiquitin ligase suppresses IFN production by down-regulating RIG-I, MDA5 and MAVS via K48-linked ubiquitination and proteasome- dependent degradation (Arimoto et al., 2007). CYLD, a deubiquitinating enzyme, functions as a negative regulator by removing these K63-linked polyubiquitin chains from RIG-I (Friedman et al., 2008). Taken together, these data implicate a coordinated network of RLH pathway ubiquitination events regulated by E3 ubiquitin ligases, DUBs and ubiquitination patterns.

4.1.2 RNF114

RNF114, also known as zinc finger protein 313 (ZNF313), is a poorly characterized RING finger protein that contains a C2HC and two C2H2 zinc-finger domains, and a C-terminal UIM (Ma et al., 2003). The human RNF114 gene is located on chromosome 20q13 and spans 17,484 bp, containing 6 exons which encodes a 228-amino acid protein (Ma et al., 2003). RNF114 was first identified by mRNA differential display between the testes of healthy adults and azoospermic patient. Two polyadenylation signals in RNF114 exon 6 were found resulting in two transcripts. The shorter transcript was expressed more abundantly in fertile adult testes than in testes of azoospermic patient suggesting that RNF114 may play a role in human spermatogenesis and male fertility

(Ma et al., 2003). In addition to testis, RNF114 is broadly expressed in many cell types,

61 including CD4+ T cells, dendritic cells and skin as well as in pancreas, kidney and spleen

(Capon et al., 2008). Previous studies have shown RNF114 to be an E3 ubiquitin ligase that can interact with both K48- and K63-linked polyubiquitin chains. Several studies have implicated RNF114 mutations as predisposing mutations in psoriasis (Capon et al.,

2008), and have suggested a positive role in mediating dsRNA responses (Bijlmakers et al., 2011). Recent studies also suggested that RNF114 may impact cell cycle regulation,

NF-κB activity and T-cell activation (Han et al., 2013; Rodriguez et al., 2014). However, the roles of RNF114 may depend on the cellular environment and cell type of its expression. Therefore an overall mechanism of action still remains unclear, and more studies are needed to clarify these questions.

62 4.2 Results:

4.2.1 RNF114 negatively impacts host innate immune responses.

4.2.1.1 RNF114 inhibits cellular dsRNA responses.

Previous studies on the RNF114 paralog RNF125 suggested that it negatively regulated the RIG-I/MDA5 innate antiviral response (Arimoto et al., 2007). Work in our lab had previously suggested that RNF114 overexpression could inhibit cellular response to double-stranded RNA (dsRNA; Li, 2007). To further characterize the physiological function of RNF114 in type I IFN regulation, we first assessed the expression profile of

RNF114 in tissues harvested from C57BL/6 mice. The steady state mRNA levels of

RNF114 in these different tissues and organs were determined by qRT-PCR. RNF114 was expressed in a variety of tissues but predominantly in the spleen and bone marrow, implicating a potential role in the immune system (Fig. 4-1A). RNF114 was also expressed across a variety of cell lines, particularly in FEMX melanoma cells, where

RNF114 levels remained largely unchanged after IFNβ treatment, suggesting that

RNF114 is not an IFN-induced gene (Fig. 4-1B). To correlate cellular dsRNA responsiveness with endogenous RNF114 expression, basal and induced ISG56 mRNA levels were measured. ISG56 is potently induced in response of type I IFNs, dsRNAs and viruses (Guo et al., 2000). Interestingly, we found that the FEMX cells did not respond to dsRNA treatment and showed decreased basal and dsRNA-induced levels of ISG56 mRNA as compared to other cell types (Fig. 4-1C). However, after knocking down

RNF114 using siRNA, ISG56 was significantly induced by dsRNA in FEMX (Fig. 4-1D).

Knocking down RNF114 using a second strategy, lentiviral shRNAs (Fig. 4-1E), also increased ISG56 and IFNβ mRNA levels in response to dsRNA treatment (Fig. 4-

63 1F).These data indicated an inverse correlation between RNF114 expression and dsRNA responsiveness.

64 A

e 100

L 80

m 60

F 40

v 20

R 0

e cle n rt en er ey n g on us ki ea le iv dn olo un B M S H Sp L Ki C L

WM9 FEMX A375 ACHN HT1080 IFN - + - + - + - + - + RNF114

actin

Hela A375 FEMX dsRNA: - + - + - +

ISG56

Actin

65 D

RNF114 mRNA ISG56 mRNA

100% 5.0

4.0 75%

3.0

50%

Expression

2.0

25%

Relative

Relative 1.0

0% 0.0

siCtl siCtl siZN F siZN F siCtl siCtl siZN F siZN F dsRN A 75ug/ml 4hr - + - + dsRN A 75ug/ml 4hr - + - +

RNF114

actin

RNF114 mRNA

1.5 Control shRNA

1 RNF114 shRNA

levels Relative 0.5

UT dsRNA UT dsRNA

66 F

ISG56 mRNA

3.5 * 3 2.5 2 N.S 1.5 1

0.5 Relative fold induction fold Relative 0 UT dsRNA UT dsRNA

IFNmRNA * 5

3 N.S 2

1 Relative fold induction fold Relative 0 UT dsRNA UT dsRNA

Control shRNA

RNF114 shRNA

67 Fig. 4-1. RNF114 inhibits cellular dsRNA responses. A) RNA was isolated from different mouse tissues and RNF114 mRNA levels were measured by qRT-PCR using mouse RNF114 primers. The relative expression levels were normalized to RNF114 mRNA levels in muscle tissue. B) RNF114 expression levels in different cell types with or without IFNβ treatment were analyzed by Western blotting using anti-RNF114 antibody. C) Cells were left untreated or treated with 100 µg /ml dsRNA for 8 h. After that time RNA was harvested and ISG56 and actin mRNA levels analyzed by RT-PCR.

(experiments were performed by Dr. Boren Lin.) D) HT1080 cells were transfected with control siRNA or RNF114 siRNA for 48 h then treated with 75 µg /ml dsRNA for 4 h.

RNA was harvested and RNF114 and ISG56 mRNA levels were analyzed by qRT-PCR and normalized to actin mRNA level. (experiments were performed by Dr. Boren Lin.) E)

RNF114 protein and mRNA levels were analyzed in control cells, control shRNA- and

RNF114 shRNA-stable transfected cells. F) Stable transfected FEMX cells (control and

RNF114 shRNA) were treated with 100 µg /ml dsRNA for 8 h then ISG56 and IFNβ mRNA levels were analyzed by qRT-PCR.

68 4.2.1.2 RNF114 suppresses RLH-dependent IFNβ production.

To identify the dsRNA response pathways inhibited by RNF114, IFNβ promoter activity was monitored by luciferase assay using an IFNβ-Luc reporter construct. Ectopic expression of either RNF114 or RNF125 resulted in a dose-dependent suppression of an

IFNβ-Luc promoter induced by RIG-I, MDA5 and MAVS (Fig. 2A-C). Consistent with that result, knocking down RNF114 by co-transfecting RNF114-targeted shRNA augmented MDA5- or MAVS-mediated IFN-Luc induction, an effect that was overcome by cotransfection of murine RNF114 that evades the human RNF114 shRNA (Fig. 2D-E).

The activity of an irrelevant, constitutive SV40 promoter was not affected by the expression of RNF114, suggesting a specific inhibitory function of RNF114 in RLH signaling (Fig. 2F). To pinpoint the site of RNF114 action on the RLH pathway, its effects on downstream components was assessed. Ectopic RNF114 overexpression did not inhibit TBK1 induced reporter activity, however, knocking down RNF114 augmented the upregulation of IFNβ promoter by TBK1 (Fig. 2G). Ectopic RNF114 expression had no significant effect on TRADD, IKKɛ or FADD induction of IFNβ or NF-κB (Fig. 2H-J).

These results indicate that RNF114 likely inhibits RLH-mediated IFNβ production at a level upstream of MAVS.

69 A B

70 RIG-I 200 MDA5

50 150

40 nduction

30 nduction 100

FoldI 20

FoldI 50 10 0 0

IFN-luc + + + + + + IFN-luc + + + + + + RIG-I - + + + + + MDA - + + + + +

RNF114 - - + ++ - - RNF1145 - - + ++ - - RNF125 - - - - + ++ RNF125 - - - - + ++

C D MAVS 700 MDA5 250

600

200 500 150 400 300 100

FoldInduction 200 FoldInduction 50 100 0 0 IFN-luc + + + + + + IFN-luc + + + + + + + MAVS - + + + + + MDA5 - + + + + + + RNF114 - - + ++ - - shRNF114 - - ++ - - ++ - RNF125 - - - - + ++ Ctrl shRNA - - - ++ - - ++ mRNF114 - - - - ++ ++ ++

70 E F MDA5 E 250 MAVS 18

200 14

12 150 10 8 100 nduction 6 FoldInduction 4

50 FoldI 2 0 0 IFN-luc + + + + + + + SV40-luc + + + + + + MAVS - + + + + + + shRNF114 - - ++ - - + - MDA5 - + + + + + Ctrl shRNA - - - ++ - - + RNF114 - - + ++ - - mRNF114 - - - - ++ ++ ++ RNF125 - - - - + ++

G H TBK1 TRADD 500 80

400

300 40 200 20

Fold Induction

Fold Induction Fold InductionFold 100

0 0 IFN-luc + + + + + + + IFN-luc + + + + + + + TBK1 - + + + + + + TRADD - + ++ +++ + ++ +++ shRNF114 - - ++ - - + - RNF114 - - - - ++ ++ ++

Ctl shRNA - - - ++ - - + mRNF114 - - - - ++ ++ ++

71 I J IKK 150 FADD 250

200 100 150

100 50

Fold Induction 50 Fold Induction

0 0 CTRL IKKi(0.1ug)+ZNF313(0.3ug) IKK - + + + + NFB-luc + + + +

RNF114 - - + ++ +++ FADD - + + + RNF114 - - + ++

72 Fig. 4-2. RNF114 inhibits RLH-mediated IFN production. 293 cells were transfected with IFN/luc, NFB/luc or SV40/luc with indicated plasmids or shRNA constructs for 24 hr followed by luciferase assay. + = 0.2 µg , ++ = 0.4 µg , +++ = 0.6 µg per 5x105 cells.

All the experiments were performed by Nichole Pfeiffer and David Velliquette.

73 4.2.2 RNF114 is an E3 ubiquitin ligase targeting MAVS for degradation.

4.2.2.1 RNF114 is an E3 ubiquitin ligase.

RNF114 contains an N-terminal RING finger domain and a C-terminal UIM, two motifs that are often found in E3 ubiquitin ligases, as well as several intervening zinc finger motifs (Giannini et al., 2008). A schematic diagram of RNF114 and the mutants used in these studies is shown in Fig. 3A. Cell-based ubiquitination assays were conducted by co-transfecting ubiquitin and His-RNF114 into HEK-293 cells, followed by nickel bead pull-down. Ubiquitin was pulled-down with His-tagged human RNF114, implicating an interaction between RNF114 and ubiquitin (Fig. 3B). Both UbcH5 and

UbcH6 E2 enzymes could conjugate Ub to mouse RNF114, as shown by using an in vitro ubiquitination assay (Fig. 3C-D). In both in vitro and cell-based ubiquitination assays,

RNF114ΔUIM (1-203), which lacks the C-terminal ubiquitin interacting motif failed to interact with ubiquitin (Fig. 3E-F). From this analysis, we concluded that the UIM of

RNF114 was required for auto-ubiquitination, and likely for transfer of ubiquitin to other substrates.

74 A

RING finger UIM C2H2 Domain RNF114FL

1 228 RNF114 ΔUIM 1 203

B C E1 - - + + + - + + E2 - 5a - 5a 5a 6 6 - mRNF114 + + + + - + + - E2 (UbcH): 2 3 5a 5b 5c 6 7 8

Ub Ub Ab Ab

RNF114 Ab D E

E1 + - + - + - + HA-Ub - - - + + + E 2 (5a) - + + - + - + FLmh - + - - + - mRNF114 - - - FL FL UIM UIM UIMmh - - + - - +

Ub Ab HA Ab

RNF114 Ab RNF114 Ab

75 Fig. 4-3. RNF114 is an E3 ubiquitin ligase. A) Schematic of RNF114 constructs used in this study. B) Purified recombinant mouse RNF114 was incubated with purified E1 and a variety of recombinant E2 for 90 min at 32 °C. Reactions were stopped by adding 4×

SDS sample buffer and were subjected to SDS polyacrylamide gel electrophoresis followed by immunoblot analysis. The membrane was probed with anti-ubiquitin antibody. C) In vitro ubiquitination assays were performed as described in the legend to

Fig. 4-3C. The membrane was probed with anti-ubiquitin antibody to detect ubiquitinated

RNF114, recombinant RNF114 was probed with anti-RNF114 antibody. D) In vitro ubiquitination assays were performed using UbcH5A as E2 ubiquitin-conjugating enzyme with full length or truncated recombinant mouse RNF114 UIM. The membrane was probed with anti-ubiquitin antibody to detect ubiquitinated RNF114, recombinant

RNF114 was probed with anti-RNF114 antibody. E) 293 cells were cotransfected with human Myc/His-tagged full length RNF114 or RNF114 UIM and HA-tagged ubiquitin for 24 h. Cells were lysed and separated by western blotting. Polyubiquitination and

RNF114 were probed with anti-HA or anti-RNF114 Ab, respectively. All the experiments were performed by David Velliquette and Dr. Boren Lin.

76 4.2.2.2 RNF114 reduced MAVS levels.

To address how RNF114 inhibits the RLH pathway, Flag-tagged MAVS or Flag- tagged MDA5 were co-transfected with control shRNA, RNF114 or RNF114 shRNA in

293 cells for 24 h. Ectopic expression of RNF114 decreased Flag-MAVS and Flag-

MDA5 protein levels while shRNA knockdown of RNF114 augmented expressions of these proteins (Fig. 4-4A). Next, we assessed whether RNF114 interacts directly or indirectly, with MAVS. Flag-MAVS or Flag-TRAF3 were co-transfected with RNF114 into 293 cells for 24 h followed by anti-Flag bead pull-down. RNF114 associated with both MAVS and TRAF3 under these conditions (Fig. 4-4B). These data suggest that

RNF114 may target MAVS and/or TRAF3 through direct or indirect binding. To further test if MAVS is a substrate of RNF114 for ubiquitination, Flag-MAVS, ubiquitin,

RNF114 or RNF125 were co-transfected into 293 cells for 24 h followed, by anti-Flag bead pull-down. RNF125, the paralog of RNF114, ubiquitinates RIG-I, MDA5 and

MAVS and mediates their proteasomal degradation to inhibit RLH-dependent IFN production. As expected, RNF125 decreased MAVS levels, and a similar effect was observed with RNF114 (Fig. 4-4C). These effects were not the result of transcriptional inhibition since the MAVS mRNA levels were unaffected. In vitro ubiquitination assay results also suggested that RNF114 could target MAVS for ubiquitination (Fig. 4-4D).

Taken together, we hypothesize that RNF114 may target MAVS for polyubiquitination mediated proteasomal degradation.

77 A

Flag-MAVS Flag-MDA5

Anti-Flag

Anti-actin

RNF114 - - - + + + Flag-MAVS - + - - + - Flag-TRAF3 - - + - - + IB:RNF114 IP:Flag

IP:Flag IB:Flag

Input IB:RNF114

78 C Flag-MAVS - + + + + + RNF114mh - - 0.2 0.6 - -

RNF125mh - - - - 0.3 0.8 Ubi + + + + + +

IP: Flag IB: Flag

Input IB: Myc

mRNA

MAVS

GAPDH

Ubi + + + + + + RNF114 - + - + + + MAVS - - ++ + ++ + + Ubi

MAVS

RNF114

79 Fig. 4-4. RNF114 decreased MAVS levels. A) Flag-MAVS or Flag-MDA5 was co- transfected with empty vector, RNF114 or RNF114 shRNA for 24 h. The expression levels of MAVS and MDA5 were analyzed by western blot. B) RNF114 was co- transfected with flag-MAVS or flag-TRAF3 for 24 h followed by flag pull-down. C)

Flag-MAVS, ubiquitin were co-transfected with RNF114 or RNF125 for 24 h followed by flag pull-down. MAVS protein and mRNA levels were analyzed by western blot and

PCR, respectively. D) in vitro ubiquitination assay was performed as described in Fig. 4-

3C using Ub, E1, UbcH5A, mouse RNF114 and MAVS. The membrane was probed with anti-ubiquitin antibody to detect ubiquitinated RNF114. Recombinant RNF114 and

MAVS were detected with anti-RNF114 and anti-MAVS antibody, respectively.

80 4.2.3 RNF114 inhibits cellular dsRNA responses in vivo.

To study the physiological function of RNF114, our laboratory generated an

RNF114 KO mouse (Lin, unpublished). Basal ISG56 mRNA levels in spleen were determined in RNF114 wild-type (WT), heterozygous and knockout (KO) animals.

ISG56 mRNA basal levels were higher in RNF114 KO mice compared to the WT mice

(Fig. 4-5A). Serum levels of IFN- were also substantially elevated in the RNF114 KO mice, indicating that RNF114 inhibits host IFN- expression in vivo (Fig. 4-5B). To assess the contribution of RNF114 to host dsRNA responses, WT and RNF114 KO mice were injected intraperitoneally with PBS or dsRNA for 8 h. As predicted by our cell based studies, RNF114 deficiency augmented dsRNA-induced IFN- serum levels and

ISG56 mRNA levels in spleen as compared with WT animals (Fig. 4-5C and D). These data suggest that RNF114 regulates basal and dsRNA-induced IFN-β production.

81 A

ISG56 mRNA ISG56

B p<0.0002

(pg/ml)

 -

Serum IFN

82 C 1000

800

(pg/ml) 600 

IFN 400

200 Serum Serum 0

RNF114 +/+ +/+ -/- -/- dsRNA - + - +

30 20

fold mRNA ISG56 10 0 RNF114 +/+ +/+ -/- -/- dsRNA - + - +

83 Fig. 4-5. RNF114 inhibits cellular dsRNA responses in vivo. A) Whole spleens from untreated animals of the indicated genotypes were assessed for ISG56 mRNA levels by qRT-PCR. Relative expression levels from individual animals is shown in the scatter plot.

B) Serum IFN- levels were determined by ELISA assay in individual animals from part

A. C) RNF114+/+ and RNF114-/- mice were injected intraperitoneally with PBS or 100

g dsRNA for 4 h. Serum levels of IFN- were determined. D) RNF114+/+ and

RNF114-/- mice were injected intraperitoneally with PBS or 100 g dsRNA for 4 h.

Splenic RNA was isolated and ISG56 mRNA levels were assessed by qRT-PCR in animals from part C. Values expressed relative to untreated control animals. All the experiments were performed by Dr. Boren Lin.

84 4.3 Discussion.

Type I IFNs play essential roles in innate immune responses against virus as well as priming the host for adaptive immune responses. As with other components of the innate immune responses, they are potently upregulated by several different pathways, including but not limited to TLR and RLH pathways. However, high or sustained levels of type I IFNs can promote inflammation and are also associated with autoimmunity

(Hooks et al., 1979; Pelegrin et al., 1998). Thus, a better understanding of the mechanisms underlying type I IFN regulation, including the details of homeostatic mechanisms within the immune system, will ultimately help us uncover potential therapeutic targets for regulating uncontrolled immune responses.

RNF114 was originally identified as a psoriasis susceptibility gene during the course of genome-wide association scans. It is expressed abundantly in disease relevant cell types, such as CD4+ T cells, dendritic cells and skin (Capon et al., 2008). However, the exact physiological and pathological mechanisms of RNF114 function are unknown.

Several studies associated RNF114 function with different processes outside of the immune system, such as in spermatogenesis, T cell activation and cell proliferation (Ma et al., 2003; Han et al., 2013; Rodriguez et al., 2014). As such that RNF114 may have a complicated, multifunctional role in a variety of signaling pathways and in different cell populations.

Although the exact function of RNF114 remains unknown, its paralogue, RNF125, has been extensively studied. RNF125 negatively regulates RIG-I and MDA5 signaling through K48-linked ubiquitination and proteasome-dependent degradation (Arimoto et al.,

2007). RNF125 serves as a negative feedback signaling molecule since it is an ISG that is

85 up-regulated by IFN. RNF125 also functions as a positive mediator of T cell activation in both Jurkat T leukemic cells and in human primary T lymphocytes (Chu et al., 2003;

Zhao et al., 2005). Recent study showed that RNF125 is downregulated in BRAF inhibitor-resistant melanomas (Kim et al., 2015). They also identified JAK1 as a target of

RNF125-dependent ubiquitination and down-regulation. Reduced RNF125 levels were associated with elevated levels of JAK1 and EGFR in BRAF inhibitor-resistant melanomas. One study showed that RNF125 mutants are associated with Overgrowth syndromes due to dysregulation of the RIG-I-MAVS, PI3K-AKT, and interferon pathways (Tenorio et al., 2014). Another study suggested that RNF125 levels were increased in IL-4-stimulated CD4+CD38- cells, resulting in suppression of HIV transcription (Shoji-Kawata et al., 2007). These findings strongly suggest that RNF114 and RNF125 may have many physiological roles which are not limited to the immune system.

Here, we focused on the role of, and the mechanism by which RNF114 regulates type I IFNs during dsRNA responses. We showed that RNF114 was ubiquitously expressed in a variety of tissues and cells and was not induced by IFNβ treatment (Fig. 4-

1A and B). Overexpression of RNF114 abrogated cellular dsRNA responses such as

ISG56 and IFNβ induction, whereas RNF114-specific siRNA/shRNA treatment augmented pathway activation. Our results suggested that RNF114 inhibited the RIG-I-,

MDA5- and MAVS-mediated pathway upstream or separately from TRADD, IKK or

FADD signaling. We also demonstrated that RNF114 is an E3 ubiquitin ligase that was auto-ubiquitinated and utilized UbcH5 and UbcH6 as an E2 (Fig. 3). Further, we showed that RNF114 associated with MAVS, although we did not determine if binding was direct

86 or indirect. RNF114 ubiquitinated MAVS both in cell and in vitro indicating that MAVS might be a substrate for RNF114 E3 activity. Exogenous RNF114 reduced MAVS and

MDA5 levels whereas knockdown of RNF114 using shRNA increased abundance of both proteins, suggesting that RNF114 may promote proteasomal degradation of MAVS and

MDA5 by poly-ubiquitination. In addition, we observed markedly elevated levels of both basal and dsRNA-induced IFNβ and ISG56 in RNF114 KO mice. Taken together, these data suggest RNF114 functions as a negative regulator of RLH pathway.

Previous studies of RNF114 showed contradicting results. One study suggested

RNF114 positively regulates cellular dsRNA responses (Bijlmakers et al., 2011). Another report showed that RNF114 down-regulates NF-κB response by stabilizing negative regulatory molecules A20 and IκBα (Rodriguez et al., 2014). However, we were unable to replicate either of these effects. The function of RNF114 may be tissue and stimuli dependent, or affected by expression levels or posttranslational modifications. Therefore, it is possible that RNF114 may possess both positive and negative regulatory functions in multiple signaling pathways, depending on the cellular context or the stimuli used.

Unlike RNF125, which has been proposed to suppress IFNβ expression after viral infection as a negative feedback mechanism, RNF114 is constitutively expressed and not upregulated by viral infection or IFNβ treatment. Therefore, we hypothesize that RNF114 down regulates MAVS as a means of maintaining low basal levels of IFNβ in the absence of pathogen in order to prevent uncontrolled inflammation. So far, however, we have not observed any autoimmune disease or any chronic inflammatory conditions in RNF114-/- mice. We also failed to observe increased viral resistance or augmented antiviral responses within RNF114-/- animals. This might be partially due to the abundance of

87 negative regulators of this pathway that compensate for RNF114 function, or to the massive upregulation of the response even in the presence of these inhibitors.

In summary, ubiquitination is a well know mechanism for targeting protein for degradation, activation and protein interactions, and is a major posttranslational modification regulating antiviral responses. Our data suggest that RNF114 is an E3 ubiquitin ligase that negatively regulates basal activity of the RLH pathway. We have demonstrated that MAVS is a potential target for RNF114-mediated polyubiquitination and degradation. This study provides more information on the potential physiological function of RNF114 in inflammation and host antiviral responses.

88 4.4 Overall Summary.

Our lab has a long-standing interest in innate immune signaling and host response to virus infections. Here, we have looked at the innate immune response from both the cell’s point of view, and the virus’ viewpoint. We demonstrated that VHSv IVb M protein potently inhibited host transcription mediated by all three RNA polymerases. Our data showed that VHSv M interfered with RNAP II promoter binding, suggesting that M might target general transcription factors. We also identified an aspartic acid residue at position 62 that was critical for M inhibitory function. Overall our data suggested that

VHSv may behave similarly to VSV by utilizing M protein to inhibit host innate immune responses by shutting down cellular transcription. With this information, we’ll conduct studies on identifying M-associated host factors to understand more clearly how M inhibits host transcription. A long-term goal of this study would be to develop an attenuated virus that can still replicate, but with reduced ability to block host antiviral signaling. If achieved, such a product might serve as an effective attenuated viral vaccine for inoculating farmed fish.

From the cell’s perspective, we reported that RNF114 is an E3 ubiquitin ligase that appears to play a role in type I IFN regulation. RNF114 inhibited RLH-dependent

IFN production and its ectopic expression reduced levels of major RLH signaling molecules, including MDA5 and MAVS, presumably by ubiquitination-mediated proteasomal degradation. In KO animal studies, we observed that RNF114-defecient mice exhibited augmented basal and induced dsRNA responsiveness. However, we did not observe any change in antiviral or inflammatory responses in those animals. Taken

89 together, we hypothesize that RNF114 maintains a low basal levels of IFN expression in the steady state to prevent unwanted inflammatory responses.

Overall, these two projects contribute to a better understanding of the need for innate immune suppression in viral pathogenesis as well as in normal cellular homeostasis. Future work will try to translate these basic research observations into applications of therapeutic interest, whether in the fish or human realm.

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