Influenza virus H5N1 non-structural protein 1 alters

interferon-α/βα/βα/βα/β signaling

By

Danlin Jia

A thesis submitted in conformity with the requirements for the degree of Master of

Science

Graduate Department of Immunology

University of Toronto

Copyright by Danlin Jia 2009

ii

ABSTRACT

Influenza virus H5N1 non-structural protein 1 alters interferon-α/βα/βα/βα/β signaling

Danlin Jia, Master of Science,

Department of Immunology, University of Toronto. 2009

Type I interferons (IFNs) function as the first line of defense against viral

infections by modulating numerous biological processes to establish an antiviral state and

influencing the activation of various immune cells. During influenza A infection, the NS1

encoded by the virus genome disrupts many cellular processes to block type I IFN

responses. We show that expression of H5N1 NS1 in HeLa cells reduces IFN-inducible

activation of STAT proteins and its subsequent binding to DNA complexes. Subsequent

analysis suggests NS1 blocks IFN signaling by inhibiting expression of type I IFN

receptor subunit, IFNAR1, as well as up-regulating SOCS1 expression. Finally, we

demonstrate that pretreatment of primary human lung tissue with IFN alfacon-1 inhibits

H5N1 viral replication by up-regulating a number of interferon-stimulated genes. The

data suggest that NS1 can directly interfere with Type I IFN signaling, and that

pretreatment with IFN can inhibit H5N1 infection in primary human lung tissue.

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ACKNOWLEDGEMENT

I would first like to express my sincere gratitude to my supervisor, Dr. Eleanor

Fish for all of her amazing guidance and support throughout the study. Her brilliance in

scientific research, sport and artistic craftsmanship has made great impact during my

graduate life, and I believe it will serve as my guidance for the future. I would like to

especially thank her for being so understanding during some of difficult time in both

research and personal life. I would like to wish her all the best in the future (where her

IFN mimetic will be the prime choice for any clinical IFN therapy).

I am also deeply grateful for my committee members Dr. Dana Philpott and Dr.

Scott Gray-Owen for their continuous support (of reference letters) throughout the study.

I wish them both all the best in the future. Furthermore, I would like to thank our

collaborator Dr. John Nicolls for providing the human sample for the viral study. In

addition, I like to thank Dr. Bing Sun and Ke Xe for providing valuable reagents for my

study.

Of course, none of this would have been possible without the rest of “Fish pond”,

and I would like to express my deep gratitude and best wishes to all of following fish

members:

To Beata: my lab mom, I thank you for all of your help and support, and I know

that your kindess will always bring me joy wherever I go. I will keep on

practicing and improving my Polish for many reasons (wink, wink!) I

wish you and Andre a life time of joy and happiness (same goes to Sam

and Smokey).

To Ramtin: my brother (from previous life, of course…), I thank you for

supporting and pushing me through all of the hurdles for the last of

couple years. Your charismatic personality has strengthened me in

ways I could never image (in a good way, of course). Your

determination in life (actually, just the part in weight lost..kidding)

will inspire me for life. I know you have something huge waiting for

you, and I wish you and Julia a lifetime of happiness and joy!

To Thomas: you have been a great mentor for me throughout the time that we

spent together. Your warm charisma and brilliance in science and

sport brings me motivation and joy. I wish you all the best and a

lifetime of happiness with Aida!

To Daniel: (this is a bit difficult, I am still “thinking” about it since I just “fainted”)

Your adventurous personality and kindness (but not your impulse

shopping habit!) will always brings me courage in life. I wish you all

(including Andrea, Megan and Joe) the best from the bottom of my

heart!

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To Carole: thank you for all the support in the past few years, I will miss you (and

Kaycee) dearly. I wish you all and your family all of best in the years

to come.

To Erin: Thank you for everything, and I will miss your headband and rest of your

fashion equipment. I know you will do good on your MCAT and your

career as a physician. I wish you and Jeff a lifetime of happiness (Tell

Jeff that I know he can make it big someday!)

To Jay: thank you for all your help (and Kimchi!) throughout the study. I wish

you all the success in the future and a lifetime of happiness with Megan,

Joshua and David.

To Olivia: you are a very talented young student with unimaginable potential (that

sounds dangerous…just kidding), believe in yourself and you will

succeed. I wish you (and Jay Chao) happiness always!

To Joanne: hey, Dude, thank you for all of your support for the past year, I wish

you all of best in your future study!

I owe my loving thanks to my family for all of their encouragement and support throught

my graduate study.

I like to gratefully thank Canadian Institute of Health Research (CIHR) and Ontario

Graduate Scholarship for Science and Technology (OGSST) for their financial support.

Toronto, Canada, May 2009

Danlin Jia

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TABLE OF CONTENTS

Chapter I

INTRODUCTION………………………………………………………………... 1-51

I.1 Types of IFN…………………………………………………………………….1

I.2 Induction of type I IFN………...……………………………………………….. 2

I.2.1 Toll-like Receptors…………………………………………………… 3

I.2.1.1 TLR signaling pathway to activation of type I IFN…………4

I.2.1.2 TLR4………………………………………………………... 5

I.2.1.3 TLR3………………………………………………………... 9

I.2.1.4 TLR7/8/9…………………………………………………… 9

I.2.2 RLH…...…………………………………………………………….... 10

I.2.2.1 RLH signaling pathway to activation of type I IFN………... 11

I.2.3 IRF in type I IFN induction…………………………………………... 12

I.3 Transcriptional regulation of type I IFN………………………………...13

I.4 Type I IFN receptors…………………………………………………………….14

I.5 Type I IFN signaling…………………………………………………………….15

I.5.1 JAK-STAT pathway………………………………………………….. 16

I.5.2 PI3’K and Akt pathway………………………………………………. 20

I.5.3 PI3’K and mammalian target of rapamycin (mTOR) pathway………. 21

I.5.4 V-crk sarcoma virus CT10 oncogene homolog(avian)-like

(CrkL)pathway……………………………………………………….. 22

I.5.5 Mitogen-activated protein kinase (MAPK) p38 pathway…………….. 22

I.6 Antiviral effectors………………………………………………………………. 23

I.6.1 dsRNA-dependent protein kinase R………………………………….. 24

I.6.2 2’-5’ oligoadenylate synthetase (2’5’OAS)…………………………...25

I.6.3 The Mx proteins……………………………………………………….26

I.7 Biological response of type I IFN……………………………………………….27

I.7.1 Antiviral……………………………………………………………….27

I.7.2 Antiproliferative and apoptosis………………………….…………… 28

I.7.3 Immunomodulation………………………………….……………….. 28

I.8 Viral evasion of type I IFNs system…………………………….……………… 29

I.9 Influenza A virus………………………………………………………….……. 30

I.9.1 Orthomyxoviridae family…………………………………………….. 30

I.9.2 Components of influenza A…………………………………………... 31

I.9.3 Influenza A replication cycle………………………………….………37

I.9.4 Genetic drift and genetic shift of influenza virus…………………….. 41

I.9.5 Influenza A repertoire and restriction……………………………….... 41

I.9.6 Clinical symptoms and treatments……………………………………. 42

I.10 NS1 and host innate immune responses……………………….……………… 43

I.10.1 Structures of NS1…………………………………………………….46

I.10.2 Functions of NS1……………………………………………………. 46

I.10.2.1 NS1 inhibits intracellular sensor RIG-I and PKR…………............ 46

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I.10.2.2 NS1 inhibits host mRNA processing and export………………….. 47

I.10.2.3 NS1 stimulates viral protein translation…………………….…….. 48

I.10.2.4 NS1 and cell survival……………………………………………… 49

I.10.3 NS1 and virulence…………………………………………………... 49

I.11 Thesis objective………………………………………………………………..51

Chapter II

MATERIALS AND METHODS……………………………………………….... 52-58

II.1 Cells, virus and reagents...…………...…………………………………………52

II.2 Transfection and virus infection………………………………………………..53

II.3 Immunoblotting and immunoprecipitation……………………………………..53

II.4 Cell sorting and flow cytometry analysis……………………………………...54

II.5 RNA extraction and cDNA synthesis…………………………………………..55

II.6 Real time-polymerase chain reaction (RT-PCR)……………………………… 57

II.7 Electrophoretic mobility shift assay…………………………………………… 57

II.8 Immunohistochemistry and Confocal microscopy……………………………..58

Chapter III

RESULTS…………………………………………………………………………. 59-82

III.1 H5N1 NS1 localizes primarily in the nucleus of HeLa cells………………… 59

III.2 H5N1 NS1 expression in HeLa inhibits IFN-inducible

STAT phosphorylation…………………………………………………......... 62

III.3 H5N1 NS1 protein expression in HeLa inhibits IFN-inducible

STAT:Sis-Inducible Element (SIE) complex formation………..…………….67

III.4 Expression of H5N1 NS1 elevates SOCS1 but not SOCS3

expression…………......................................................................................... 70

III.5 H5N1 NS1 expression in HeLa leads to reduction in surface IFNAR1

but not surface IFNAR2………………………………………………………70

III.6 Both H5N1 NS1 expression in HeLa and H5N1 influenza A infection in

primary human lung tissue down-regulates IFNAR1 but not IFNAR2

mRNA expression………………………………………………………......... 76

III.7 Reduction of IFNAR1 but not IFNAR2 gene expression is not

result of their differential mRNA halflife…..………………………………... 79

III.8 Pretreatment with type I IFN up-regulates ISGs and inhibits H5N1

influenza A replication in primary human lung cells…………....................... 82

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Chapter IV

DISCUSSION……………………………………………………………………... 85-93

Chapter V

FUTURE DIRECTIONS..………………………………………………………... 94-95

Chapter VI

REFERENCES……………………………………………………………………. 96-127

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LIST OF FIGURES

INTRODUCTIONS

Figure I.1 Induction of type I IFNs………………………………………… 7

Figure I.2 Type I IFNs signaling……………………………………………17

Figure I.3 Schematic representation of influenza A virus…………………. 32

Figure I.4 Replication of influenza A virus………………………………... 39

Figure I.5 Interaction between influenza A NS1 and host molecules………44

RESULTS

Figure III.1 H5N1 NS1 localizes predominantly in the nucleus of HeLa

cells………………………………………………...…………. 60

Figure III.2 H5N1 NS1 expression inhibits IFN-inducible STAT

phosphorylation……………………………………..………… 64-66

Figure III.3 H5N1 NS1 protein expression inhibits IFN-inducible

STAT:SIE complex formation…..…………………………… 68

Figure III.4 Expression of H5N1 NS1 reduces surface IFNAR1

but not IFNAR2 expression….………………………………..72

Figure III.5 Expression of H5N1 NS1 reduces IFNAR1 but not

IFNAR2 mRNA expression……………………….................... 77

Figure III.6 NS1-mediated reduction of IFNAR1 but not IFNAR2

gene expression is not reflective of differential mRNA

half-life………………………………………….…………….. 73

Figure III.7 Expression of H5N1 NS1 elevates SOCS1 but not SOCS3

Expression…………………………………………………….. 80

Figure III.8 IFN-alfacon-1 inhibits H5N1 influenza A replication and

induces up-regulation of ISGs in primary human lung cells…. 83

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LIST OF ABBREVIATIONS

2-5A 2’-5’ oligoadenylate

2’5’OAS 2’-5’oligoadenylate synthetase

4E-BP1 eIF4E-binding protein

AP-1 activation protein-1

APRE acute phase response element

BRG Brahma-related gene

BAF BRG-BRM-associated factor

CARD caspase-recruitment domains

CD cluster of differentiation

CID central interactive domain

CIS cytokine-inducible SH2-containing

CPSF cleavage and polyadenylation specificity factor

CrkL V-crk sarcoma virus CT10 oncogene homolog (avian)-like

c-Src cellular sarcoma

DBD DNA binding domain

DD death domain

DsRNA double stranded ribonucleic acid

eIF4E eukaryotic translation initiation factor 4E

ERK extracellular signal regulated kinases

FADD Fas-associated death domain

FBN fibronectin

GAS γ-activating sequence

GCN5 general-control-amino-acid synthesis 5

GEF guanine exchange factor

HA hemagglutinin

HAT histone acetyl transferases

HIV human immunodeficiency virus

HTLV human T-cell leukemia virus

IKK IκB kinase

IL interleukin

IκB inhibitor of NFκB

IRS insulin receptor substrate

ISG interferon-stimulated gene

JAK Janus kinase

LGP2 likely ortholog of mouse D11lgp2

LPS lipopolysaccharide

LRR leucine-rich repeat

LZ leucine zipper

M matrix

MAL MyD88 adaptor-like

MAK mitogen-activated protein kinase

MAKK MAK kinase

MAKKK MAKK kinase

x

MDA5 melanoma differentiation antigen 5

mTOR mammalian target of rapamycin

Mx Orthomyxovirus resistance

NA neuraminidase

ND nuclear domain

NFκB nuclear factor-kappa B

NK natural killer

NLS nuclear localization signal

NP nucleoprotein

NS nonstructural

OPN osteopontin

PAB polyadenylate binding protein

PAMP pathogen-associated molecular pattern

pDC plasmacytoid dendritic cell

PI3K phosphatidylinositol 3-kinase

PIAS protein inhibitor of activated STAT

PKB protein kinase B

PKC protein kinase C

PML NB promyelocytic leukemia protein nuclear body

PRD positive regulatory domain

PRE prolactin response element

PRR pattern recognition receptor

PTP protein tyrosine phosphatase

RD repressor domain

RIG-I retinoic-inducible gene I

RLH RIG-like helicase

S6K S6 kinase

SARM sterile α- and armadillo-motif-containing protein

SIE sis-inducible element

SH src-homology

SOC suppressor of cytokine signaling

ssRNA single stranded RNA

STAT signal transducer and activator of transcription

SUMO small ubiquitin-related modifier

TAB TAK1-binding protein

TAD transcriptional activation domain

TAK TGF-β-activated kinase

TANK TRAF-family-member-associated NFκB activator

TBK1 TANK-binding kinase 1

TCP T cell-PTP

THOV Thogotovirus

TIR toll/interleukin 1 receptor

TIRAP TIR-associated protein

TLR Toll-like receptor

TNF tumor necrosis factor

TOP terminal oligopyrimidine

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TRAF tumor necrosis factor (TNF) receptor associated factor

TRAM TRIF-related adaptor molecule

TRIF TIR-domain-containing adaptor protein-inducing IFN-β

tRNA transfer RNA

Ubc13 ubiquitin conjugating enzyme 13

Uev1A ubiquitin-conjugating E2 enzyme variant 1A

1

CHAPTER I

INTRODUCTION

IFN was discovered by Isaac and Lindenmann in 1957 as a secreted substance that

confers antiviral activity against influenza infection (Isaacs and Lindenmann, 1957). The

discovery of IFNs paved the way for understanding other class II cytokines and their

receptors. IFNs are pleiotropic cytokines that are produced in response to viral challenge,

and function to protect the host against infection via the transcriptionl and translational

induction of a series of proteins that interfere with different stages in the replicative cycle

of viruses (Samuel, 1991). In addition, IFNs activate a number of immune cells, thereby

invoking the clearance of virus (Le Bon and Tough, 2002).

I.1 Types of IFN

There are three types of IFN, each characterized by their distinct cognate

receptors. The Type I IFNs are comprised of different subtypes including: multiple IFN-

α subtypes (14 human, 11 mouse), IFN-β, IFN-ε, IFN-ω, IFN-δ and IFN-τ (Hardy et al.,

2004). Type I IFNs can be produced by most cell types and bind as monomers to the two

subunits of the type I IFN alpha receptor, IFNAR1 and IFNAR2 (de Weerd et al., 2007).

IFN-γ is the sole Type II IFN, is functionally active as a dimer, and binds with high

affinity to its cognate receptor that is comprised of the two subunits IFNGR1 and

IFNGR2 (Soh et al., 1994; Soh et al., 1993). Unlike type I IFNs, IFN-γ is only produced

by few cell types, including natural killer (NK) cells, CD4+ T helper 1 (Th1) cells, and

dendritic cells (DC). The third type of IFN has 3 members, IFNλ1, IFNλ2 and IFNλ3,

also known as interleukin-29 (IL-29), IL-28A, and IL-28B, respectively (Kotenko et al.,

2

2003; Sheppard et al., 2003). Type III IFNs bind with high affinity to the IFN lambda

receptor subunits, IFNLR1 and IFNLR2. Type I and type II IFNs have distinct and

overlapping signaling pathways as well as biological activities. Current evidence suggests

that type III IFNs share many of the signaling and responses of type I IFNs (Dumoutier et

al., 2003). Different type I IFNs exhibits differential affinity for its receptor, but currently

the IFN that has the highest affinity for type I IFN receptors belong to the recombinant

IFN named IFN-alfacon-1 (Ozes et al., 1992). This IFN was generated by comparing the

most frequent occurring amino acid among endogenous type I IFNs (Pfeffer, 1997).

I.2 Induction of type I IFNs

Type I IFNs are rapidly induced when viral or bacterial derived factors, also

known as pathogen-associated molecular patterns (PAMPs) interact with cellular pattern

recognition receptors (PRRs) (Medzhitov and Janeway, 1997). PRRs include members of

the toll-like receptor (TLR) family, cytosolic sensors like retinoic acid-inducible gene I

(RIG-I)-like helicase (RLH), nucleotide-oligomerization domain (NOD)-like receptors

(NLRs) and the DNA-dependent activator of IRFs, DAI. PRR activation leads to

downstream signalling cascades and the production of type I IFNs and/or other pro-

inflammatory cytokines (Kawai and Akira, 2006). During the late 1980s, Charles

Janeway hypothesized that PRRs would be capable of detecting a broad range of

infectious agents, and subsequently induce appropriate immune responses to combat

infection. This concept led to the subsequent identification and characterization of

PAMPs, shared across different microbial families (Medzhitov and Janeway, 1997).

These conserved moieties are not only “foreign” to the host, enabling discrimination from

3

self, but also represent molecules that are intolerant to extensive mutations because of

their critical roles in the clearance of microbes.

I.2.1 Toll-like Receptors

Toll, a protein involved in Drosopila embryogenesis, was found to play a critical

role in the immune response to fungus infection (Medzhitov et al., 1997). This led to the

subsequent discovery and characterization of its mammalian homologue and related

family members. There are currently 10 human TLRs and 13 murine TLRs (Takeda et al.,

2003). Each TLR acts alone or in combination with other TLRs to detect unique PAMPs.

TLR4 was first shown to be involved in the recognition of lipopolysaccharide (LPS), a

component of Gram-negative bacterial cell walls (Poltorak et al., 1998). TLR1, in

combination with TLR2, recognizes triacyl lipopeptides, whereas when complexed with

TLR6, TLR2 can also bind diacyl lipopeptide (Hajjar et al., 2001; Shimizu et al., 2005).

TLR3 is the receptor for double stranded RNA (dsRNA) (Alexopoulou et al., 2001).

TLR5 is activated in the presence of bacterial flagellin (Hayashi et al., 2001). TLRs7/8

recognize ssRNA, whereas TLR9 binds to unmethylated dsDNA (Heil et al., 2004;

Hemmi et al., 2002; Takeshita et al., 2001). TLRs are also differentially located in cells,

with TLR1, 2, 4, 5 and 6 found on the plasma membrane and TLR3, 7, 8 and 9 located

inside late endosomes or lysosomes (Kawai and Akira, 2005). TLRs are type I

transmemebrane proteins, each composed of extracellular leucine-rich repeats (LRRs)

followed by one or two cysteine-rich regions which are involved in ligand binding. TLRs

have short transmembrane domains connected to a cytoplasmic toll/interleukin 1 receptor

4

homology (TIR) domain, which functions in the recruitment of downstream adaptor and

signaling components in response to ligand recognition (Akira, 2004).

I.2.1.1 TLR signaling activates type I IFN production

TLR-ligand interactions lead to changes in receptor conformation, allowing the

cytoplasmic TIR domain to interact with several downstream TIR-containing adaptor

proteins, including MyD88, MyD88 adaptor-like (MAL or TIR-associated protein

[TIRAP]), TIR-domain-containing adaptor protein-inducing IFN-β (TRIF or TIR-

domain-containing molecule 1 [TICAM1]), TRIF-related adaptor molecule (TRAM) and

sterile α- and armadillo-motif-containing protein (SARM) (O'Neill and Bowie, 2007).

Differential usage of these adaptors can result in different downstream responses, which

are distinguished by two pathways: the MyD88-dependent pathway and the MyD88-

independent pathway. All but TLR3 can activate the MyD88-dependent pathway,

whereas only TLR3 and TLR4 can signal through the MyD88-independent cascade

(Kawai and Akira, 2005; Oshiumi et al., 2003; Yamamoto et al., 2002). Notably, only

TLR3, 4, 7, 8 and 9 activation leads to the production of type I IFNs (Figure I.1). Current

evidence suggests stimulation of TLR1, 2, 5 and 6 leads to signaling through the classical

MyD88-dependent nuclear factor-kappa B (NFκB) pathway, resulting in the production

of pro-inflammatory cytokines such as interleukin-6 (IL-6) and tumor necrosis factor α

(TNF-α) but not IFNs (Bas et al., 2008; Fisette et al., 2003; Wang et al., 2001b; Zeng et

al., 2006).

5

I.2.1.2 TLR4

TLR4 is expressed on the cell surface, and its extracellular portion associates with

MD-2, a secreted polymeric protein required for the oligomerization of TLR4 in response

to LPS. MD-2 is also involved in the glycosylation of TLR4, a necessary process for its

translocation to the plasma membrane (Shimazu et al., 1999). In addition, TLR4-MD2

can associate with CD14, a co-receptor for LPS (Jiang et al., 2005). Upon ligand

activation, the TLR4-MD2-CD14 complex recruits both adaptor MAL and TRAM,

initiating both MyD88-dependent and MyD88-independent signaling, respectively (Seya

et al., 2005). On one hand, in the absence of MyD88, TRAM can interact with the adaptor

TRIF, which in turn leads to association with the tumor necrosis factor (TNF) receptor

associated factor 6 (TRAF6) (Hoebe et al., 2003; Yamamoto et al., 2003b). TRAF6

subsequently activates TRAF-family-member-associated NFκB activator (TANK)-

binding kinase 1 (TBK1) and the non-canonical inhibitor of NFκB (IκB) kinase (IKKi)

(Hemmi et al., 2004; Sato et al., 2003). TBK1 will phosphorylate interferon regulatory

factor 3 (IRF3), allowing it to dimerize and translocate to the nucleus, where it can induce

the transcription of IFN-β (Sakaguchi et al., 2003). TLR4 can also interact with MyD88

via the TIR domain through the adaptor MAL, which leads to the downstream activation

of the interleukin-1 receptor (IL-1R) associated kinase -4 and -1 (IRAK-4 and IRAK-1),

TRAF6 and TGF-β-activated kinase 1 (TAK1) (Dong et al., 2006; Fitzgerald et al., 2001;

Loiarro et al., 2007; Muzio et al., 1997; Suzuki et al., 2002). Though understanding the

precise relationship between these kinases requires ongoing investigation, current

evidence suggests binding of MyD88 brings IRAK4, IRAK1 and TRAF6 together at the

receptor complex. IRAK4 subsequently phosphorylates IRAK1, leading to its activation

6

and autophosphorylation (Cheng et al., 2007; Kollewe et al., 2004; Li et al., 2002).

Hyperphosphorylated IRAK1 dissociates from MyD88 to form a cytoplasmic complex

with TRAF6 via interactions between their death domains (DD)(Ahmad et al., 2007; Qian

et al., 2001). TRAF6 in turn interacts with a multiprotein complex including TAK1-

TAK1-binding protein 1 (TAB1)-TAB2 and ubiquitin-conjugation enzyme 13 (Ubc13)

and ubiquitin-conjugating E2 enzyme variant 1A(Uev1A)(Deng et al., 2000; Takaesu et

al., 2000). TAK1 activation leads to phosphorylation of the IKK complex (including

IKKα, IKKβ and scaffolding IKKγ), which subsequently leads to the phosphorylation of

IκB and its degradation(Lee et al., 2000; Takaesu et al., 2003; Wang et al., 2001a). This

in turn facilitates the release and translocation of NFκB into the nucleus. TAK1 can also

phosphorylate mitogen-activated protein (MAP) kinases including extracellular signal-

regulated kinase (ERK) 1 and 2, c-Jun N-terminal kinases (JNKs) and p38, leading to the

activation of activation protein-1 (AP-1), which can function cooperatively with either

IRF3 or NFκB to induce the expression of type I IFNs or proinflammatory cytokines,

respectively (Thiefes et al., 2005; Wang et al., 2001a; Yang et al., 2004).

7

Figure I.1 Induction of type I IFNs. Viral derived-factors or PAMP (red arrow) activate

PRRs like TLRs (TLR3,4,7,8,9) and cytoplasmic sensors (RIG-I, MDA5, DAI) to induce

transcriptional activation of type I IFNs mediated by IRF3 and IRF7.

8

9

I.2.1.3 TLR3

TLR3 is predominantly located in endosomes and phagosomes, the exceptions

being TLR3 expression on the cell surface of epithelial and natural killer (NK) cells

(Matsumoto et al., 2003; Schmidt et al., 2004; Xie et al., 2007). TLR3 recognizes dsRNA,

a common replicative intermediate during virus infection. Evidence suggests acidification

of endosomes is the first step toward initiating TLR3 activation (de Bouteiller et al.,

2005). Downstream signaling from TLR3 is mediated in a MyD88-independent fashion,

with recruitment of TRIF first to its TIR domain, followed by aggregation with TRAF3

(Oganesyan et al., 2006; Yamamoto et al., 2003a). TRAF3 in turn activates kinases

TBK1 and IKKi, leading to the subsequent phosphorylation of IRF3, which dimerizes

and translocates to the nucleus to activate IFN-β gene expression (Hacker et al., 2006;

Yoneyama et al., 1998). In addition, TLR3-dsRNA interactions induce phosphorylation

of TLR3 cytoplasmic tyrosine residues, which then function as docking sites for

phosphatidylinositol 3-kinase (PI3K) (Johnsen et al., 2006; Sarkar et al., 2004).

Activation of PI3K in response to dsRNA is required for the complete phosphorylation of

IRF3 (Dong et al., 2008). Furthermore, tyrosine kinase cellular sarcoma (c-Src) kinase

can also be activated and associate with TLR3 in response to dsRNA, though its role in

downstream signaling is yet to be determined (Johnsen et al., 2006).

I.2.1.4 TLR7/8/9

Human plasmacytoid dendritic cells (pDC) lack TLR3 and TLR4, yet express

high levels of TLR7 and TLR9, whose activation leads to robust type I IFN production in

response to virus-derived single stranded RNA (ssRNA) or unmethylated CpG DNA

10

motifs, respectively (Hornung et al., 2002). TLR7, 8 and 9 invoke the MyD88-dependent

signaling pathway (as described above) to induce both type I IFNs and proinflammtory

cytokines. In contrast to TLR3, the induction of type I IFNs from these TLRs does not

require the presence of IRF3 (Honda et al., 2005). Following ligand activation, MyD88,

IRAK4, IRAK1 and TRAF6 recruitment to the receptor results in activation of

downstream TAK1. This leads to the activation and subsequent nuclear translocation of

NFκB and AP-1. In addition, TRAF3 can also aggregate with MyD88, IRAK1, IKKα

and precursor of osteopontin (OPN) to activate IRF7 (Honda et al., 2004; Kawai et al.,

2004; Oganesyan et al., 2006). Phosphorylated IRF7 can subsequently dimerize and

translocate to the nucleus, where it interacts with the promoters in the IFN-α and IFN-β

genes to activate their transcription (Kawai et al., 2004; Yeow et al., 2000). For both

TLR8 and TLR7, ligand recognition overlaps, thereby confounding the identification of

distinguishing signaling pathways. TLR8 may function as a negative regulator for TLR7

and TLR9 (Wang et al., 2006b).

I.2.2 RLH

The induction of type I IFNs in the absence of TLRs led to the identification of

TLR-independent viral sensors. The RNA helicase RIG-I was the first cytoplasmic

receptor identified capable of sensing dsRNA and activating type I IFN production

(Yoneyama et al., 2004). RIG-I is comprised of a C-terminal DExD/H box RNA helicase

domain that interacts with dsRNA in an ATP-dependent manner, two N-terminal caspase-

recruitment domains (CARDs) which can interact with other CARD-containing proteins,

and a repressor domain (RD), that has been shown to suppress signaling in the resting

11

state(Saito et al., 2007). Two other members of the RIG-I-like helicases (RLH) have been

described: melanoma differentiation antigen 5 (MDA5) and likely ortholog of mouse

D11lgp2 (LGP2). Like RIG-I, MDA5 contains two CARD-like domains and a helicase

domain. In contrast, LGP2 only possesses the helicase domain and does not contain a

CARD-like domain. LGP2 may function as a negative regulator of RIG-I and MDA5

(Yoneyama et al., 2005). Notably, RIG-I and MDA5 exhibit some specificity in the viral

PAMPs they recognize. Gene targeting studies have shown that RIG-I is critical for the

detection of paramyxoviruses, vesicular stomatitis virus (VSV) and influenza viruses,

whereas MDA5 is vital for the recognition of picornaviruses (Childs et al., 2007; Kato et

al., 2006; Loo et al., 2008).

I.2.2.1 Type I IFN production mediated by RLH signaling

Upon binding to target RNA, RIG-I first undergoes ubiquitination of its CARD

domain. This ubiquitin E3 ligase tripartite motif 25 (TRIM25) catalyzed reaction is

critical for efficient downstream signalling (Gack et al., 2007). In contrast, ubiquitination

by another protein, RNF125, targets RIG-I for degradation, suggesting RIG-I-mediated

signaling is tightly regulated through differential ubiquitination (Arimoto et al., 2007).

Subsequent to ligand recognition, RIG-I undergoes a conformational change and self-

association, which in turn leads to binding with its downstream adaptor IFN-β promoter

stimulator 1 (IPS-1, also known as MAVS, VISA or CARDIF) via CARD-CARD

interactions. IPS-1 is comprised of an N-terminal CARD domain, a proline rich region

and a hydrophobic transmembranous (TM) region at the C-terminus (Kawai et al., 2005).

The TM region anchors IPS-1 to the outer membrane of the mitochondrion, required for

12

subsequent signaling (Seth et al., 2005). Activated IPS-1 can associate with TRAF3 to

activate downstream kinases TBK1 and IKKi, which in turn can phosphorylate IRF3 and

IRF7 to induce type I IFN gene expression (Kawai et al., 2005; Xu et al., 2005). IPS-1

can also interact with Fas-associated death domain-containing protein (FADD), a

mediator in death receptor signalling (Kawai et al., 2005). FADD associates with

caspase-8 and caspase-10, leading to their cleavage and the activation of downstream

NFκB, resulting in the production of pro-inflammatory cytokines (Kreuz et al., 2004).

However, the precise mechanism leading to NFκB activation from caspase cleavage

remains to be determined. In addition to the RLH family, there are other cytoplasmic

sensors that can respond to PAMPs. DNA-dependent activator of IRFs (DAI) was

recently identified to induce an IFN response when stimulated with dsDNA, and its

presence seems to be important for detecting DNA viruses (Takaoka et al., 2007).

I.2.3 IRF in type I IFN induction

The IRF family of proteins is comprised of nine members (Taniguchi et al., 2001).

IRF3 and IRF7 play key roles in mediating type I IFN gene expression. IRFs possess a

conserved N-terminal DNA binding domain (DBD) with five tryptophan repeats (Harada

et al., 1994; Taniguchi et al., 2001; Veals et al., 1992). Crystal structure studies have

identified 5’-AANNGAAA-3’ as the consensus base sequence recognized by IRFs (Fujii

et al., 1999; Tanaka et al., 1993). Binding to this sequence leads to changes in DNA

structure that may allow cooperative binding of other transcription factors like AP-1 and

NFκB to nearby target sequences. Under normal conditions, both IRF3 and IRF7 reside

in the cytoplasm in inactive forms. Upon PRR activation, IRF3 and IRF7 undergo serine

13

phosphorylation mediated by activated TBK1 and IKKi (as described above), and form

either homodimeric or heterodimeric complexes (Fitzgerald et al., 2003; Hemmi et al.,

2004; Sharma et al., 2003). These dimers translocate to the nucleus and complex with

other co-activators to target specific gene elements in the promoters of type I IFNs and

other cytokines, thereby activating transcription (Marie et al., 2000). In contrast to IRF3,

which plays a major role in regulating IFN-β expression, IRF7 can activate gene

expression of the IFN-αs and IFN-β. IRF7 is generally less abundant in cells than IRF3,

with the exception of cells of lymphoid origin, particularly plasmacytoid dendritic cells

(pDC) (Izaguirre et al., 2003). The expression of IRF7 can be up-regulated in response to

IFN-β stimulation, which in turn can act to induce gene expression for the IFN-αs.

Maintenance of this positive feedback loop is thought to require the ongoing presence of

viral factors, since IRF7 has a short half-life due to its susceptibility to ubiquitin-

mediated degradation (Negishi et al., 2005). Other IRF members such as IRF1 and IRF5

have also been implicated in type I IFN production, though gene targeting studies suggest

that both are dispensable for normal IFN expression (Reis et al., 1994; Schoenemeyer et

al., 2005).

I.3 Transcriptional regulation of type I IFNs

Transcription of the IFN-β gene requires the assembly of the transcriptional

complex, also known as the enhanceosome, at the enhancer region upstream of the IFN-β

gene transcription start site. The enhancer region of the IFN-β gene contains four positive

regulatory domains (PRDs I, II, III and IV), whereas genes for the IFN-αs contain PRD-

I- and PRD-III-like elements (PRD-LE) (Kim and Maniatis, 1997; Ryals et al., 1985).

14

Activated IRFs recognize PRD-I and PRD-III, whereas PRD-II is targeted by AP-1 and

PRD-IV by NFκB. These activated transcriptional factors associate with the high-

mobility group protein I (Y) (HMG I [Y]) to form the enhanceosome (Kim and Maniatis,

1997). The enhanceosome subsequently recruits histone acetyl transferases (HATs),

including general-control-control-amino-acid synthesis 5 (GCN5) and CREB-binding

protein (CBP/p300) to catalyze histone (H3 and H4) acetylation. This modification

results in engagement of Brahma-related gene (BRG)-Brahma (BRM)-associated factor

(BAF) complex, which leads to spatial alternation in the nucleosome and facilitates the

binding of the RNA polymerase complex to the start site of transcription (Agalioti et al.,

2000).

I.4 Type I IFN receptors

Following PRR activation and the subsequent induction of expression for Type I

IFNs, these secreted proteins function in both autocrine and paracrine ways to influence

multiple cellular functions to establish an antiviral state. For Type I IFNs to exert their

influence in cells, the absolute requirement is that these cells express the two

transmembrane receptor subunits of the IFN receptor, IFNAR: IFNAR1 and IFNAR2c.

These receptors were identified through a series of genetic cloning and IFN sensitivity

reconstitution assays using somatic cell hybrids (Langer and Pestka, 1988). In contrast to

IFNAR1, human IFNAR2 has three isoforms as a result of alternative splicing and

differential usage of exons and polyadenylation. IFNAR2c represents the isoform

comprised of an extracellular domain, a transmembrane region and a cytoplasmic domain

(Lutfalla et al., 1995). In contrast, IFNAR2a is a soluble receptor. IFNAR2b is similar to

15

IFNAR2c, but lacks a cytoplasmic domain (Novick et al., 1995). Interestingly, mice do

not have the IFNAR2b isoform, instead there are two soluble IFNAR2a murine variants,

arising from differential splicing (Owczarek et al., 1997). Binding to and engagement of

both IFNAR1 and IFNAR2c are required for productive type I IFN signalling. The type I

IFNs bind with higher affinity to IFNAR2 compared with IFNAR1, and IFNAR2c is

considered the primary binding receptor subunit (Jaks et al., 2007). The extracellular

portion of IFNAR1 is comprised of four domains named SD 1-4, with each domain

containing a fibronectin (FBN) III-like motif. SDs 1-3 appear to contribute to ligand

binding, whereas SD4 appears critical for the formation of the receptor complex (Ghislain

et al., 1994; Lamken et al., 2005). On the other hand, all isoforms of IFNAR2 contain two

FBN-like domains configuring as an immunoglobulin-like folding structure (Chill et al.,

2003; Kumaran et al., 2007; Runkel et al., 2000). The binding of type I IFNs to these

cognate receptors leads to activation of receptor-associated Janus kinases (JAKs), which

in turn phosphorylate tyrosine residues in the intracellular domains of each receptor

subunit, leading to the recruitment of signaling effectors, their phosphorylation-activation

and subsequent activation of signaling cascades. In contrast to the non-signaling receptor

subunit IFNAR2b, soluble IFNAR2a is capable of acting as both activator and inhibitor

in the context of IFN signaling, yet more studies are required to elucidate its precise

function (Fernandez-Botran, 1991; Han et al., 2001; Hardy et al., 2001).

I.5 Type I IFN signaling

Binding of type I IFNs to IFNAR leads to receptor aggregation and activation of

receptor-associated kinases: JAKs. The JAK family is comprised of JAK1, 2, 3 and

16

TYK2, each containing a protein kinase domain at their carboxyl-terminus and five other

domains that make up the N-terminus (Darnell et al., 1994; Stark et al., 1998). TYK2 and

JAK1 interact with the cytoplasmic domains of IFNAR1 and IFNAR2, respectively.

Activation of these kinases leads to phosphorylation of cytoplasmic tyrosine residues in

the IFNARs, thereby generating docking sites for src-homology 2 (SH2)-containing

signaling molecules (Platanias and Colamonici, 1992). The recruitment and

phosphorylation of downstream molecules initiates a series of signaling pathways,

leading to both transcriptional and translational activation (Figure I.2).

I.5.1 JAK-STAT pathway

Proteins that play a prominent role in mediating the transcriptional activation of

type I IFNs are the signal transducers and activators of transcription (STAT) proteins

(Figure I.2). There are seven members in this family: STAT1, STAT2, STAT3, STAT4,

STAT5a, STAT5b and STAT6 (Copeland et al., 1995; Fu et al., 1992; Schindler et al.,

1992). STAT proteins, by their name, can relay signals from a wide spectrum of

cytokine-receptor complexes, including IFN-, interleukin- and growth factor- receptor

compexes (Bromberg, 2001; Schindler, 2002). Each STAT has a dimerization domain at

the N-terminus, a coil-coil domain, a DNA binding domain, a linker domain, an SH2

domain and a carboxyl transcriptiononal activation domain (TAD) (Levy and Darnell,

2002). STATs were first thought to exist as monomers in the cytoplasm in the absence of

receptor stimulation, but recent evidence indicates that many STATs form dimers and

shuttle between the cytoplasm and nucleus even without cytokine-receptor activation

17

Figure I.2 Type I IFN signaling. Binding of type I IFNs to their cognate receptors leads

to activation of receptor-associated kinases Jak1 and Tyk2, and initiates both

transcriptional and translational activation to establish an antiviral response

18

.

19

(Koster and Hauser, 1999; Mao et al., 2005). In the presence of type I IFNs, STAT

dimers are recruited to phosphorylated IFNARs through their SH2 domains and become

phosphorylated by JAKs (Shuai et al., 1993; Silvennoinen et al., 1993). This tyrosine-

phosphorylation leads to STAT homo- or hetero-dimerization, mediated by the binding of

the SH2 domain of one STAT to the phospho-tyrosine of another (Becker et al., 1998;

Chen et al., 1998; Mao et al., 2005). Activated STAT dimers subsequently enter into the

nucleus via importin-mediated translocation, and target specific DNA elements to

activate the expression of IFN-stimulated genes (ISGs) (Friedman et al., 1984; Larner et

al., 1984; McBride et al., 2002). A number of STAT complexes are formed in response to

type I IFN stimulation, with IFN stimulated gene factor 3 (ISGF3) being one of the best

characterized complexes (Fu et al., 1990; Levy et al., 1989). ISGF3 is comprised of

STAT1, STAT2 and IRF9/p48. Nuclear translocation enables ISGF3 to target gene

promoters containing an IFN stimulated response element (ISRE), to activate the

expression of ISGs including ISG15, 6-16 and ISG54 (Darnell et al., 1994). Though

complexes like STAT2:2:IRF9 and STAT2:6:IRF9 can also target ISREs in the

promoters of genes, they exhibit a much lower binding affinity when compared to ISGF3.

In addition, other activated complexes like STAT1:1, STAT1:3, STAT3:3 and STAT1:2

target another group of ISGs whose promoters contain the consensus γ-activating

sequence (GAS) element (Brierley and Fish, 2005a; Ghislain and Fish, 1996; Vinkemeier

et al., 1996). Variants of GAS elements include acute phase response element (APRE),

sis-inducible element (SIE) and prolactin response element (PRE) (Decker et al., 1997).

Activation of ISG expression functions to inhibit virus infection mediated by different

antiviral proteins. STAT signaling is negatively regulated by members of the cytokine-

20

inducible SH2-containing protein (CIS)/suppressor of cytokine signaling (SOC) family,

protein inhibitors of activated STAT (PIAS) and other protein tyrosine phosphatases

(PTPs) (Liu et al., 2004; Song and Shuai, 1998). SOC family members contain a variable

N-terminus, an SH2 domain, and a SOCS box domain at the carboxy terminus (Hilton et

al., 1998; Krebs and Hilton, 2000). Though there are eight SOC family members, only

SOC1, SOC3 and CIS are well characterized for their role in regulating type I IFN

signaling. SOC1 inhibits IFN signaling through direct physical interaction with JAK,

whereas SOC3 and CIS interact with the phosphorylated receptor to hinder the

recruitment and phosphorylation of downstream mediators like STAT proteins

(Matsumoto et al., 1999; Yasukawa et al., 1999). In contrast to SOCS whose expression

are rapidly induced post cytokine stimulation, PIAS proteins are constitutively expressed

and interact directly with activated STATs to block their DNA-binding activity (Liu et al.,

1998; Starr et al., 1997). Other phosphatases like SH2 domain-containing PTP-2 (SHP-2)

and T cell-protein tyrosine phosphatase (Tc-PTP) 45 (TCP45) have also been shown to

inactivate STATs through dephosphorylation (Simoncic et al., 2002; Wang et al., 2006a).

I.5.2 PI3K and Akt pathway

Type I IFNs can also trigger the activation of the PI3K pathway to modulate

cellular translation and survival. Upstream effectors in this signaling cascade are the

adaptor protein insulin receptor substrates (IRSs). IRSs were originally identified as

critical mediators of insulin signalling (Myers et al., 1993; White, 1998). They contain

residues that can undergo phosphorylation to become docking sites for downstream SH2-

containing signaling moieties (Platanias et al., 1996). IFN-α/β-receptor interactions result

21

in IRS-1 and IRS-2 phosphorylation by JAK1, leading to the interaction between IRS-1

and the regulatory subunit of PI3K, p85, through its SH2 domain (Burfoot et al., 1997).

Phosphorylation of p85 activates the catalytic subunit of PI3K, p110, which can in turn

activate one of its downstream effectors, protein kinase B (PKB)/Akt, through the

generation of phosphatidylinositol 3, 4, 5-triphosphate (PIP3) (Manning and Cantley,

2007; Uddin et al., 1997). Activation of PKB influences numerous cellular processes

including both proliferation and survival (Manning and Cantley, 2007; Martelli et al.,

2007).

I.5.3 PI3K and mammalian target of rapamycin (mTOR) pathway

IFN-inducible activation of PI3K can regulate protein translation by modulating

the activity of the mTOR pathway (Manning and Cantley, 2003). Indeed, the Akt

pathway has a role in mRNA translation (Kaur et al., 2008). Activated mTOR signals

through two major downstream effectors, namely, the S6 kinase (S6K) and the eukaryotic

translation initiation factor 4E (eIF4E)-binding protein 1(4E-BP1) to modulate protein

translation (Hara et al., 1997; von Manteuffel et al., 1997). Through a physical

association, 4E-BP1 blocks the interaction of eIF4E with other translation initiation

factors, thereby preventing cap-dependent translation (Scheper et al., 1992).

Phosphorylation of 4E-BP1 by mTOR relieves its association with eIF4E, thereby

allowing its association with eIF4G and other co-factors to initiate translation (Hara et al.,

1997). mTOR also influences translation through the activation of S6K. Activated S6K

phosphorylates the ribosomal protein S6 to increase the translation of 5’-terminal

22

oligopyrimidine (TOP) mRNAs, which include ribosomal proteins and elongation factors

(Jefferies et al., 1997; Tang et al., 2001).

I.5.4 V-crk sarcoma virus CT10 oncogene homolog (avian)-like (CrkL) pathway

CrkL belongs to the Crk family of proteins which include CrkI and CrkII. Crk

proteins contain both SH2 and SH3 domains and function as adaptors in cytokine

signalling (Mayer et al., 1988). IFN-α/β binding to IFNAR results in the activation of

TYK2, leading to the phosphorylation of Casitas B-lineage lymphoma (CBL), an adaptor

protein that associates constitutively with TYK2 (Uddin et al., 1996). Phosphorylated

CBL acts to recruit CrkL via its SH2-binding domain. CrkL subsequently recruits a range

of downstream effectors including C3G, a guanine exchange factor (GEF) for Rap-1

(Feller et al., 1995; Reedquist et al., 1996; Sattler et al., 1996; Tanaka et al., 1994). Rap-1,

identified first as a tumor suppressor gene, inhibits the activity of the small GTPase Ras,

to hinder cellular proliferation (Cook et al., 1993). CrkL can form a complex with

phosphorylated STAT5 through its SH2 domain to activate the expression of GAS-

containing genes (Fish et al., 1999).

I.5.5 Mitogen-activated protein kinase (MAPK) p38 pathway

The three major MAPK families are extracellular signal regulated kinases (ERKs),

JNKs, and the p38 MAP kinases (Schaeffer and Weber, 1999). These serine-threonine

kinases respond to various stimuli and coordinate numerous signaling cascades to

generate appropriate cellular responses (Kyriakis and Avruch, 1996). The p38 MAP

kinase family is comprised of four p38 isoforms (α, β, γ and δ) that can be activated in

23

response to stress (radiation, heat shock and hyperosmolarity), and cytokines such as

interleukin-1 (IL-1), transforming growth factor-β (TGF-β) and tumor necrosis factor-α

(TNF-α) (Jiang et al., 1996; Jiang et al., 1997; Lechner et al., 1996; Lee et al., 1994;

Raingeaud et al., 1995). Evidence from various studies suggests that upon IFN

stimulation, Vav, a GEF, is phosphorylated by TYK2, and will activate the small G-

protein Ras-related C3 botulinum toxin substrate 1 (Rac1) (Crespo et al., 1997; Platanias

and Sweet, 1994). Rac1 activation initiates a series of downstream phosphorylation

cascades involving MAPK kinase kinase (MAPKKK) and MAPK kinase (MAPKK),

including MKK3/4/6 to activate p38 MAP kinases (Salojin et al., 1999). p38 MAPKs

target other molecules such as MapKapK-2 and MapKapK-3, two serine kinases which

have important roles in mediating the antiviral effect of type I IFNs (Mayer et al., 2001;

Uddin et al., 1999). Although there have been reports that p38 MAPK may phosphorylate

serine residues on STAT1, protein kinase C-δ (PKC-δ) is primarily responsible for the

IFN-α/β inducible serine phosphorylation of STATs (Kovarik et al., 1999; Uddin et al.,

2002). Phosphorylation of serine 727 on STAT1 and STAT3 is required to achieve their

full transcriptional activation (Wen et al., 1995; Zhu et al., 1997).

I.6 Antiviral effectors

Type I IFNs are pleiotropic cytokines that influence numerous cellular processes.

One of the major downstream responses is to generate effectors that inhibit the replication

of pathogens. IFN-inducible transcriptional activation of ISGs and IFN-inducible

regulation of translational events lead to a defined set of proteins being expressed that can

exert inhibitory effects at different stages of viral replication to achieve an antiviral effect.

24

I.6.1 dsRNA-dependent protein kinase R

dsRNA-dependent protein kinase R (PKR) is one of the well characterized ISG

products that participates in both IFN-inducible antiviral and antiproliferative responses.

PKR is a serine-threonine kinase comprised of a kinase domain and two dsRNA binding

domains (dsRBD) (Patel and Sen, 1992). PKR is activated by dsRNA, a common

replicative intermediate during virus infection. Binding of dsRNA to PKR leads to PKR

dimerization and subsequent autophosphorylation (Galabru and Hovanessian, 1987). This

activated PKR acts on downstream effectors to modulate both translation and

transcription. One of the well known targets of PKR is eIF2α, a factor that plays a critical

role during the initiation of translation (Rhoads, 1993; Williams, 1999). eIF2 is made up

of three subunits: α, β and γ, that together function to promote the guanine trisphosphate

(GTP)-dependent delivery of Met-transfer RNA (tRNA) to the 40S ribosome during

protein synthesis (Hershey, 1991). Next, GTP is hydrolyzed and allows eIF2 to dissociate

from the initiation complex (Majumdar and Maitra, 2005). Subsequently, eIF2B, a GEF,

will recycle the inactive eIF2-GDP back to its active form. However, activation of PKR

leads to phosphorylation of eIF2α, which leads to an increase in its affinity for eIF2B.

The sequestration of eIF2B results in the inhibition of Met-tRNA delivery and thereby

prevents the initiation of translation of both viral and cellular mRNAs (Hershey, 1991).

In addition, PKR can modulate the activity of transcription factors in response to

dsRNA. PKR plays a role in the phosphorylation of serine residues in STAT1 and STAT3

(Ramana et al., 2000). Abrogation of this PKR-mediated serine phosphorylation leads to

loss of function of these STATs (Deb et al., 2001; Lee et al., 2005). PKR has also been

25

reported to participate in the activation of NFκB, acting on its upstream IKK kinase

complex (Gil et al., 1999, 2000). Activated IKKs lead to the phosphorylation of IκB,

thereby promoting translocation of activated NFκB into the nucleus. Through the

regulation of both transcriptional and translational pathways, PKR is able to regulate

cellular apoptosis, growth and differentiation.

I.6.2 2’-5’ oligoadenylate synthetases (2’5’OAS)

Similar to PKR, 2’5’OAS are IFN-inducible proteins that act in a dsRNA-

dependent manner (Zhou et al., 1993). There are three members in the OAS family

including OAS1, OAS2 and OAS3. These proteins contain a conserved domain known as

the 2’-5’OAS unit that corresponds to the first 346 amino acids (Hovnanian et al., 1998).

Though OAS proteins lack the classical binding site for dsRNA, data from crystal

structures and mutagenesis studies suggest that there is a conserved motif for ligand

recognition (Hartmann et al., 2003; Rebouillat et al., 1999). In the presence of dsRNA,

2’5’OAS is activated and begins to synthesize 2’-5’ oligonucleotides of adenylate (2-5A)

of various sizes, dependent on the specific OAS. 2-5A is a ligand for RNase L, a

riboendonuclease comprised of nine ankyrin repeats, a number of protein-kinase like

motifs and an RNase domain (Zhou et al., 1993). Binding of 2-5A transforms the inactive

monomeric RNase L into its activated dimeric conformation, which functions to cleave

single-stranded RNA at the 3’ side of the UpAp or UpUp regions (Dong and Silverman,

1995; Dong et al., 1994; Tanaka et al., 2004). This process antagonizes viral replication:

(1) cleavage of viral genomic RNA (Li et al., 1998), (2) degradation of viral mRNA, (3),

because RNase L cleaves both viral and host mRNA species, the availability of host

26

proteins required for viral replication will be affected (Banerjee et al., 2000; Smith et al.,

2005). In addition, cleavage of single-strand RNA can generate short dsRNAs that can

serve as ligands for cytosolic sensors like RIG-I and MDA5 (Malathi et al., 2007).

Activation of these cytosolic PRRs will lead to the additional production of type I IFNs

and enhance antiviral responses. Lastly, RNase L activation has also been associated with

the induction of apoptosis through cytochrome c and caspase 3-mediated events, invoking

another mechanism for inhibiting viral replication (Castelli et al., 1997; Rusch et al.,

2000).

I.6.3 The Mx proteins

Orthomyxovirus resistance (Mx) proteins were originally identified as IFN-

sensitive factors in a mouse strain (A2G) that exhibited higher resistance to influenza

virus infection compared to other mouse strains (Horisberger et al., 1983; Lindenmann,

1964). Members of the Mx family belong to the large GTPase family (Staeheli et al.,

1993) and contain a GTPase domain at the N-terminus, a central interactive domain

(CID), and a leucine zipper (LZ) motif at the C-terminus (Haller et al., 2007). The

cellular localization of Mx proteins varies among the different isoforms and is also

species-dependent. The mouse Mx1 protein resides in the nucleus, as a result of a nuclear

localization signal (NLS) in its C-terminus, and its mechanism of antiviral action differs

from cytoplasmic Mx proteins such as MxA (human) and Mx2 (rodent) (Staeheli and

Haller, 1985; Staeheli et al., 1986; Staeheli et al., 1993). Inside the nucleus, Mx1 resides

in a subnuclear partition known as the promyelocytic leukemia protein nuclear body

(PML NB) or also known as nuclear domain 10 (ND10), and interacts with molecules

27

such as Sp100, Daxx and factors of small ubiquitin-related modifier-1 (SUMO-1)

(Chelbi-Alix et al., 1995; Engelhardt et al., 2001; Trost et al., 2000). The precise antiviral

mechanism of action of Mx1 during viral infection remains unknown. Interestingly,

experiments with Thogotovirus (THOV), a member of the orthomyoxvirus family,

suggest that the cytoplasmic MxA protein can sequester viral nucleocapsids to prevent

their translocation into the nucleus, where viral replication and virion pre-assembly takes

place (Kochs and Haller, 1999a, b). This activity does not seem to require the GTPase

activity of MxA, but the oligomerization of MxA seems to be important for both ligand

recognition and protein stability (Haller and Kochs, 2002).

Distinct from those described above, other IFN-inducible proteins have been

shown to participate in mediating type I IFNs responses, e.g. TRAIL, viperin, p21, p200

family members, caspases, though for some, their precise mechanisms of action requires

ongoing investigation (Brierley and Fish, 2005b).

I.7 Biological response of type I IFNs

I.7.1 Antiviral

The signature response of type I IFNs is the induction of cellular resistance to

viral infection. By modulating different cellular signaling cascades, type I IFNs can

inhibit virus infection by targeting different stage(s) of viral replication. IFN-α has the

ability to inhibit the entry of VSV through secreting antiviral factor(s) that remain to be

identified (Basu et al., 2006). Hepatitis virus entry is also blocked by IFNs, mediated by

down-regulation of the viral cell surface receptor, SR-BII (Murao et al., 2008). The

28

effector functions of IFN-inducible PKR, Mx and 2’5’OAS/RNase L discussed above,

can cooperate to inhibit viral replication and viral protein synthesis. Studies with human

immunodeficiency virus (HIV) and human T-cell leukemia virus type 1 (HTLV-1)

suggest IFNs can also exert their inhibitory effects at the viral assembly stage (Dianzani

et al., 1998; Feng et al., 2003; Oka et al., 1990). Furthermore, IFN-inducible ISG15

expression can inhibit the budding of Ebola virus through disrupting the ubiquitination of

its VP40 protein (Okumura et al., 2008).

I.7.2 Antiproliferative effects and apoptosis

IFN-α was the first cytokine used clinically for anti-tumour therapy, based on

potent antiproliferative activity. Currently, recombinant IFN-αs are used clinically for

certain haematological malignancies (Kirkwood and Ernstoff, 1984). Type I IFNs

downregulate the expression of oncogenes such as c-myc, and up-regulate negative cell

cycle regulators including IRF1 and JUND (Jia et al., 2007; Knight et al., 1985;

Papageorgiou et al., 2007). There are many ISGs that code for pro-apoptotic factors to

regulate cell survival. Studies in cells of different lineages indicate that type I IFNs can

up-regulate Fas (CD95), caspases, members of regulator of apoptosis family, and other

factors that function to stimulate or sensitize target cells to apoptosis (Juang et al., 2004;

Selleri et al., 1997). Certainly, induction of cell death is also an effective mechanism

against viral infection.

29

I.7.3 Immunomodulation

Type I IFNs modulate functions of effector cells from both the innate and

adaptive immune system (Brierley & Fish, JICR 2002). IFNs will promote the survival of

memory T cells via IL-15 stimulation (Rogge et al., 1998). IFNs induce B cell maturation

and influence immunoglobulin (Ig) class switching. NK cells are activated by type I IFNs

to increase effector functions (Biron et al., 1999). Expressions of chemokines and

chemokine receptors are also regulated by type I IFNs, allowing differential trafficking of

immune effectors to sites of inflammation (Salazar-Mather et al., 2002).

I.8 Viral evasion of IFN antiviral effects

Given the critical role of type I IFNs as a first line of defence against infection, it

is not surprising that many viruses have evolved ways to target this system as a means to

increase their replication efficiency. Viral-mediated inhibition of type I IFNs can be

generalized into three categories, including disruption of IFN induction, disruption of

IFN-inducible signaling and disruption of IFN-mediated effector functions. The non-

structural protein 3/4A (NS3/4A) of Hepatitis C virus (HCV) is a serine protease that can

cleave both IPS-1 and TRIF. Disruption of these adaptor proteins blocks RIG-I- and

TLR3-mediated IFN induction (Li et al., 2005; Meylan et al., 2005). Paramyoxviruses

such as simian virus 5 (SV5), mumps virus and parainfluenza virus, all express a V

protein which is capable of blocking MDA5-mediated signaling through direct physical

interactions(Andrejeva et al., 2004). Furthermore, V protein inhibits IFN signaling by

targeting STAT proteins for proteasome degradation (Didcock et al., 1999). The poxvirus

vaccinia virus (VACV) encodes a soluble viral receptor IFNAR homologue, capable of

30

blocking IFN signalling (Colamonici et al., 1995). The soluble viral receptor sequesters

extracellular IFN to prevent its interaction with cell surface expressed IFNAR. VACV

also encodes other viral proteins that target both TLRs and their adaptors to block the

activation of IRF3 and NFκB, thereby inhibiting IFN production (Bowie et al., 2000).

I.9 Influenza A virus

Influenza virus is one of the leading infectious pathogens that threaten public

health. The mortality rate of annual outbreaks ranges from a quarter to half a million

around the globe, and brings morbidity to 5% -15%. Infection by influenza A virus

strains accounts for the majority of severe outbreaks. The severity of influenza A virus

infections was well demonstrated in the devastating 1918 Spanish flu outbreak, in which

an estimated 20 to 25 million lives were lost around the world (Hampton, 2004). In 1957

and 1968 there were two other influenza virus infection pandemics and between one to

four million lives were lost during each outbreak (Palese, 2004). Given this history, the

recent emergence of a highly virulent avian influenza A H5N1 infection in humans has

raised serious public health concerns of another potential pandemic. Interestingly, the

word ‘influenza’ is derived from the Italian word “ influence,” when in the mid 1700s

people believed celestial stars were somehow affecting people with disease (DeLacy,

1993; Heilman and La Montagne, 1990).

I.9.1 Orthomyxoviridae family

Influenza viruses A, B, C, and thogotoviruses together constitute the

orthomyxoviridae family. The types of influenza viruses can be distinguished based on

31

the antigenic properties of their nucleoprotein (NP) and matrix protein 1 (M1) (Cheung

and Poon, 2007). Influenza A can be further categorized into different subtypes based on

the antigenic properties of two of its surface glycoproteins: hemagglutinin (HA) and

neuraminidase (NA). There are currently 14 HA and 9 NA that are used to designate

influenza A strains in the form of HxNx (Fouchier et al., 2005; Laver et al., 1984).

I.9.2 Components of influenza A

Influenza A virus is an enveloped virus whose genome is comprised of eight

segments of negative sensed ssRNA that encode ten different proteins (Figure I.3) (Palese,

1977). The virion contains three surface proteins, including HA, NA and M2 (Cheung

and Poon, 2007). The cytoplasmic side of the viral envelope associates with M1 (Ruigrok

et al., 1989). The eight segments of the ssRNA genome are wrapped around NP proteins,

associated with the viral RNA polymerase and non-structural protein 2 (NS2) (Lamb and

Choppin, 1983; Richardson and Akkina, 1991; Yasuda et al., 1993).

Influenza A viral RNA polymerase

Influenza A encodes an RNA polymerase that is responsible for its transcription

and replication. This RNA-dependent RNA polymerase is comprised of three subunits:

PB2, PB1 and PA. PB2 is encoded by the first viral gene segment and functions to bind

and cleave the 5’cap structures of the host mRNA through its endonucleolytic activity

(Blaas et al., 1982; Ulmanen et al., 1983; Ulmanen et al., 1981). The cleaved

oligonucleotide is utilized both as a primer for the initiation of viral transcription and as a

32

recognition site for translation. PB1 is encoded by the second segment, and catalyzes the

RNA-dependent RNA polymerization during viral transcription and viral genome

33

Figure I.3 Schematic representation of influenza A virus.

34

35

replication (Biswas and Nayak, 1994; Kobayashi et al., 1996). The RNA polymerase

contains conserved motifs shared by other RNA-dependent RNA polymerases, as well as

domains that interact with PB2 and PA (Digard et al., 1989). The precise function of PA

is not yet clear, but evidence suggests it plays a role in both transcription and

endonucleolytic cleavage of the viral polymerase during viral replication (Fodor et al.,

2002; Zurcher et al., 1996).

Hemagglutinin (HA)

HA is an integral protein expressed on the surface of the influenza virion. It is

responsible for viral entry, binding to the sialic acid-containing host cell receptors. Sialic

acids belong to the nine carbon monosaccharide family detected on the terminus of

glycolipids and glycoproteins. HA, depending on its subtype, can recognize sialic acid in

either a sialic α2-3 galactose linkage or a sialic α2-6 galactose linkage (Connor et al.,

1994; Vines et al., 1998). The activity of HA requires several post-translational

modifications, including cleavage of its signal peptide, glycosylation, palmitylation and

trypsin-like protease mediated cleavage to generate HA1 and HA2 (Horimoto et al.,

1994). HA forms a homotrimer on the virion surface with HA1 forming a head and HA2

being the stalk of the monomer (Wilson et al., 1981). Due to the low fidelity viral RNA

polymerase, the influenza genome is constantly subjected to random mutations, thus

leading to the generation of different HA subtypes. There are fourteen antigenically

distinct HAs, designated HA 1-14, thereby affecting the nature of seasonal vaccines and a

consideration for any pandemic vaccine.

36

Nucleoprotein (NP)

The influenza NP is a 56kD protein that is abundantly expressed in the virion. It

possesses an N-terminal domain that interacts with viral RNA, motifs required for

oligomerization, and domains that interact with the viral RNA polymerase (Albo et al.,

1995; Biswas et al., 1998; Kobayashi et al., 1994; Mena et al., 1999). The NP associates

with vRNA and viral polymerase to form the viral ribonucleoprotein complex (vRNP)

inside the virion. The NP also contains a nuclear localization signal (NLS), which may be

responsible for the nuclear transport of vRNP during infection (O'Neill et al., 1995;

Whittaker et al., 1996). In addition, the expression level of NP may regulate the switch

between viral mRNA synthesis and viral genome replication by interacting with viral

RNA polymerase (Shapiro and Krug, 1988).

Neuraminidase (NA)

The influenza NA is the second major surface protein of the influenza virion. It is

expressed as a homotetramer and functions to cleave terminal sialic acid from

glycoproteins or glycolipids (Hausmann et al., 1997). This activity is important for the

budding of the virion, since the presence of sialic acid at the budding site can hinder viral

egress via interactions with surface HA molecules (Palese et al., 1974). Like HA, NA is

also glycosylated and subject to constant mutations and pressure from the immune system.

There are currently nine NA subtypes that are antigenically distinct, designated NA 1-9.

37

Matrix proteins (M1 and M2)

Both M1 and M2 are encoded by segment seven in the influenza virus genome.

M1 represents the most abundant viral protein found in the infectious particle. It

functions as the major structural protein that encapsulates the vRNP complex inside the

viral membrane (Ruigrok et al., 2000; Ye et al., 1999). There is evidence that M1

participates in several aspects of viral replication, including inhibition of viral replication,

regulation of vRNP transport and viral assembly (Martin and Helenius, 1991; Watanabe

et al., 1996). M2 is an integral membrane protein, generated from segment seven via

alternate splicing and forms a homotetrameric complex on both the host cell and virion

surface (Holsinger and Lamb, 1991; Lamb et al., 1985). The M2 tetramer forms the

proton channel and can regulate the pH inside the virion (Ciampor et al., 1992). The

reduction of pH via M2 is critical for the dissociation of M1 from vRNP, a process that is

essential for subsequent viral transcription (Bui et al., 1996).

Nonstructural proteins (NS1 and NS2)

The last segment of the influenza genome also encodes two viral proteins as a

result of alternate splicing. The colinear transcript encodes NS1, the sole viral protein that

is absent in the infectious particle but expressed abundantly as a dimer early during

infection(Nemeroff et al., 1995). NS1 resides predominately in the nucleus and is well

known for its ability to inhibit type I IFNs responses (Garcia-Sastre et al., 1998). NS1

interacts with other host factors to modulate transcription, translation, survival and

apoptosis, which will be discussed in more detail below. A splice variant of the same

gene segment for NS1, NS2, is present in the infectious virion in small quantities (Ward

38

et al., 1995). Current evidence suggests that NS2 can stimulate viral replication via a

hitherto unknown mechanism (Odagiri et al., 1994). NS2 also contains a nuclear export

signal that is important for nuclear export of vRNPs at the late stage of infection

(Neumann et al., 2000).

I.9.3 Influenza A replication cycle

Upon infection, influenza A virus first binds to sialic acid-containing

glycoproteins on the host cell mediated by HA molecules (Figure I.4). The virus then

enters the cell via clathrin-mediated endocytosis. Once inside the endosome, the

reduction of pH leads to a conformational change that unmasks the fusion peptide of the

HA molecule that is required for joining the viral envelope and the endosomal membrane

(Ciampor et al., 1992; Skehel et al., 1982). Simultaneously, the M2 ion channel starts

importing hydrogen ions into the interior of the virion, leading to the dissociation of the

M1 oligomer and dissociation of the M1- viral RNA polymerase complex, thereby

facilitating the release of vRNPs into the cytoplasm (Bui et al., 1996). vRNPs in turn are

transported into the nucleus and primary transcription is initiated. The viral RNA

polymerase starts to remove 5’cap structures from host mRNA and incorporates these

onto the viral mRNA to facilitate transcription and subsequent translation (Blaas et al.,

1982; Ulmanen et al., 1983; Ulmanen et al., 1981). Viral proteins produced early during

infection include the NP, NS1, and the viral RNA polymerase complex (PB1, PB2 and

PA) (Laine et al., 1982; Shapiro et al., 1987). These all contain an NLS and are

subsequently transported back into the nucleus upon synthesis. Subsequently, NS1 will

act to block host mRNA splicing and export, interacting with factors of the splicing

39

machinery and those of the nuclear export complexes (Qiu and Krug, 1994; Qiu et al.,

1995). The retention of host mRNA has been proposed as a mechanism to increase the

time during which the viral polymerase can act to remove 5’cap structures from the host

mRNA. As infection progresses, depending on the expression of NP, newly synthesized

viral RNA polymerase can produce additional viral mRNAs or mediate replication of the

viral genome using viral mRNA as template. Interestingly, inhibition of host RNA

polymerase has been shown to hinder viral replication, though the precise mechanism by

which this occurs is not clear. Nonetheless, this suggests that the activity of the host

RNA polymerase is involved in influenza virus replication (Engelhardt and Fodor, 2006).

As infection progresses, viral translation shifts to produce structural proteins such as HA,

NA and M1 (Laine et al., 1982; Shapiro et al., 1987). HA, NA and M2 undergo several

post-translational modifications and localize to the plasma membrane (Hughey et al.,

1992; Jones et al., 1985; Roth et al., 1983). Meanwhile, in the nucleus NP starts to

complex with viral genome segments and viral RNA polymerase to form vRNPs. vRNPs

subsequently associate with M1 to initiate the pre-assembly of the virion (Martin and

Helenius, 1991; Watanabe et al., 1996). NS2 has also been reported to interact with

vRNPs and the M1 complex, but its role in viral assembly remains unclear. M1 proteins

of pre-assembled virions in turn interact with the cytoplasmic domain of HA and NA to

initiate viral budding from the cell plasma membrane (Ali et al., 2000). However, the

influenza A virus particle requires cleavage of HA into HA1 and HA2 to become

infectious, and this is thought to be mediated by intracellular proteases and proteases in

the respiratory tract of the host (Bottcher et al., 2006; Horimoto et al., 1994).

40

Figure I.4 Replication of influenza A virus. The virion enters the cell via endocytosis,

and subsequently fuses with the endosomal membrane to release its viral

ribonucleoprotein complex into the nucleus. Once inside the nucleus, the viral genome is

first transcribed to produce viral proteins that are needed to facilitate its genome

replication. Finally, the pre-assembled viral genome associates with structural proteins at

the cell membrane to initiate the release of virions.

41

42

I.9.4 Influenza virus genetic drift and genetic shift

Two major mechanisms contribute to the generation of novel antigenically

distinct influenza virus strains: genetic drift and genetic shift (Webster et al., 1992). The

former refers to genetic mutations created by the low fidelity viral RNA polymerase,

resulting in substitutions, insertions and deletions in any segment of the viral genome.

Each replication cycle has the potential to generate a number of variants. Although these

mutations may result in defective virus and therefore be eliminated, other mutations may

confer a selective advantage to the virus and therefore be retained. Genetic shift refers to

the reassortment of segments of the viral genome when two or more different strains of

influenza virus infect the same cell. When that happens, random incorporation of genome

segments during viral assembly can lead to the generation of novel viral strains.

I.9.5 Influenza A virus host restriction

Influenza A viruses can infect a broad range of species including human, bird, pig

and horse. Aquatic birds are the primary reservoir for influenza viruses, in which all of

the subtypes can be found (Hinshaw et al., 1980). Generally, birds are infected in the

gastrointestinal tract (GI) without producing any symptoms of disease, and virions are

excreted in the feces (Webster et al., 1978). The specific avian species that is targeted by

a particular influenza A virus is determined largely by the specificity of the viral HA and

linkage conformation of the sialic acid of the host. The HA from human influenza A

viruses preferentially binds to sialic acid α2-6 galactose linkages, which are

predominantly expressed in the upper respiratory tract. Avian and equine influenza A

viruses usually contain HAs that preferentially bind to sialic acid α2-3 galactose linkages

43

(Connor et al., 1994; Vines et al., 1998). Notably, avian influenza virus H5N1 will bind

sialic acid α2-3 galactose sites in human lower respiratory tract tissues and recent data

indicate that the respiratory tract in humans expresses both α2-3 and α2-6 linkages

(Nicholls ref). Interestingly, pigs also express both sialic acid α2-3/6 galactose linkages

and can be infected by both human and avian influenza A viruses (Kundin, 1970;

Pensaert et al., 1981; Scholtissek et al., 1983). Accordingly, public health organizations

have raised considerable concern that the potential re-assortment between human and

avian influenza virus strains may occur in pigs, and may lead to the emergence of

aggressively virulent avian-derived human tropic influenza A viruses.

I.9.6 Influenza A virus infection: clinical symptoms and treatments

The earliest symptoms of respiratory influenza A virus infection in humans are

associated with sore throat, cough, running nose and fever. Headache, chills, sweats and

muscle soreness also occur. Patients infected with avian influenza H5N1 virus exhibit

high fever, shortness of breath and gastro-intestinal disorders, such as diarrhoea. Infection

like H5N1 progresses rapidly to cause respiratory distress syndrome, renal dysfunction

and multi-organ failure, and may ultimately lead to death. Currently, there are two major

classes of drugs available for treating influenza virus infections (Thanh et al., 2008). The

first includes members of the adamantine family (amantadine, rimandtadine), developed

to inhibit the M2 ion channel to block the release of vRNP into the cytoplasm (Davies et

al., 1964; Wang et al., 1993). The second family of inhibitors that include oseltamivir and

zanamavir, target the NA protein and function to prevent viral budding (De Clercq, 2004;

Moscona, 2005). However, as a result of continuous genetic mutation, there are emerging

44

an increasing number of influenza A virus isolates that exhibit resistance to these

inhibitors, indicating the need for alternative antiviral therapies against influenza A virus

infections (Bright et al., 2005; Bright et al., 2006).

I.10 NS1 and host innate immune responses

Similar to many other viruses that have developed mechanisms to evade an IFN

response, influenza viruses have evolved to target this system. Interestingly, unlike

vaccinia virus which targets several different components of the IFN system via different

viral proteins, influenza A virus can effectively block various aspects of an IFN response

through a single protein – NS1. NS1 derived its name from the fact that it is not present in

the infectious virion but is expressed early during infection, i.e. a non-structural protein.

It is a multifunctional protein that interacts with numerous host-derived molecules

including nucleotides and proteins (Figure I.5).

45

Figure I.5 Interaction between NS1 and host molecules. NS1 can block host mRNA

processing by interfering with splicing, 3’ end processing and their export. It can also

enhance viral protein translation and cell survival, and inhibit an IFN response.

46

47

I.10.1 Structure of NS1

NS1 possesses a dsRNA binding domain (1-73) at its N-terminus that is

responsible for interacting with both viral and host ribonucleotides, and an effector

domain that comprises the rest of NS1 (Qian et al., 1995). The effector domain contains

numerous motifs that interact with a number of host proteins. NS1 is functionally active

as a dimer. Structural analysis suggests that the N-terminal domain of NS1 comprises

three α-helices that form a symmetrical six-helical fold when NS1 dimerizes (Chien et al.,

2004). Additional experiments revealed that the N-terminal domain interacts solely with

the canonical A-form of dsRNA (Chien et al., 2004). The effector domain is made up of

seven β-strands and three α-helices that are stabilized through a series of hydrophobic

interactions. This segment contains three well-characterized domains that interact with

eIF4GI, the 30kD subunit of cleavage and polyadenylation specificity factor (CPSF30)

and poly(A) binding protein II (PABII) (Burgui et al., 2003). Recent studies indicate that

this region also interacts with the p85 subunit of PI3K and CrkL, as well as members of

nuclear export complexes (Hale et al., 2008; Satterly et al., 2007).

I.10.2 Functions of NS1

I.10.2.1 NS1 inhibits intracellular sensors RIG-I and PKR

As mentioned, NS1 inhibits an IFN response by multiple mechanisms. NS1 will

disrupt the induction of IFNs by inhibiting the intracellular sensor RIG-I, mediated by

direct physical interactions (Guo et al., 2007). RIG-I plays a critical role in detecting

ssRNA during influenza A virus infection. Its activation leads to association with the

48

downstream adaptor IPS-1, and eventually leads to the phosphorylation of IRF3 and

subsequent transcriptional activation of IFN-β (Pichlmair et al., 2006; Yoneyama and

Fujita, 2007). Both IRF3 translocation and NFκB activation have been shown to be

impaired in the presence of NS1 upon dsRNA stimulation, which in turn blocks the

induction of both proinflammatory cytokines and IFNs (Donelan et al., 2004; Wang et al.,

2000). NS1 can also associate with PKR, though it remains unclear whether this

interaction is dependent on the presence of dsRNA. This association blocks PKR-

mediated phosphorylation of eIF2α and prevents translation arrest in the host during viral

infection (Lu et al., 1995).

I.10.2.2 NS1 inhibits host mRNA processing and export

Inside the nucleus, NS1 functions to disrupt post-transcriptional processing of

host mRNAs at multiple stages, including splicing, polyadenylation and nuclear export.

Pre-mRNA splicing is carried out by the splicesome, which involves a series of base pair

interactions between the transcript and a series of small nuclear RNAs (snRNAs)

including U6 snRNA (Berget and Robberson, 1986; Pironcheva et al., 1988). NS1 can

associate with U6 snRNA via an interaction between its dsRNA binding domain and the

stem loop region of U6 snRNA (Qiu et al., 1995). Binding of U6 snRNA by NS1 blocks

the subsequent interactions with other snRNAs and disrupts the catalysis of pre-mRNA

splicing. Studies have shown that transcription of the IFN-ß gene is not affected by NS1,

but that there is an accumulation of its pre-mRNA transcript in the presence of NS1 (Qiu

et al., 1995). Aside from disrupting splicing, the effector domain of NS1 can also block

the polyadenylation of host mRNA, via interactions with CPSF30 and PABII (Burgui et

49

al., 2003; Chen et al., 1999; Nemeroff et al., 1998). The generation of mature host mRNA

involves CPSF-mediated endonucleolytic cleavage of the 3’ end followed by addition of

a series of adenylates. This 3’ end processing is required for mRNA export and prolongs

mRNA half-life from the activity of endonucleases (Wang and Kiledjian, 2000).

Following 3’ cleavage, PABII functions to catalyze the elongation of the polyadenylate

tail. By binding to CPSF and PABII, NS1 suppresses the generation of mature host

mRNAs. Recently, NS1 has been shown to form inhibitory complexes with the mRNA

export machinery, including NXF1/TAP, p15/NXT, Rae1/mrnp41 and E1B-AP5 (Satterly

et al., 2007). Though the functional consequences of these interactions remains unclear,

the outcome is to block host mRNA export during infection. NS1 inhibits the expression

of a number of genes including pro-inflammatory cytokines such as TNF-α, IL-6, IL-1β,

IL18 and macrophage inflammatory protein-1 alpha (MIP-1α), and as well as ISGs such

as the IFNαs, PKR, and 2’5’OAS (Stasakova et al., 2005). However, gene expression

microarray analysis and other studies have indicated that inhibition of host gene

expression by NS1 is selective and does not affect the export of viral mRNAs. This

selectivity may relate to the ability of NS1 to recognize the 5’ untranslated region (UTR)

of viral mRNA (Chen and Krug, 2000).

I.10.2.3 NS1 stimulates viral protein translation

Aside from its ability to inhibit the induction of IFNs-α/β, NS1 can modulate

other cellular pathways during infection to support viral replication. NS1 can interact

with the translation initiation factor eIF4GI of the translational machinery via its effector

domain. This interaction in conjunction with the ability of NS1 to recognize 5’ UTR of

50

viral mRNAs have been proposed as mechanisms by which the virus is able to

specifically enhance the translation of viral proteins during infection (Enami et al., 1994;

Salvatore et al., 2002).

I.10.2.4 NS1 and cell survival

In vitro, NS1 expression can induce apoptosis of a number of cell types, including

Madin-Darby Canine Kidney (MDCK) and lung epithelial carcinoma A549 (Schultz-

Cherry et al., 2001; Zhang et al., 2007) cells. Caspases and p53 activation are elevated in

the presence of NS1. Interestingly, recent studies revealed that NS1 can activate PI3K via

an interaction with the regulatory subunit of PI3K, p85, through a putative SH2-binding

domain. Activation of PI3K by NS1 in turn leads to the downstream activation of Akt

(Ehrhardt et al., 2007; Shin et al., 2007). Activated Akt can regulate numerous cellular

processes including cell survival. Indeed, the presence of NS1 has been shown to delay

cell apoptosis during infection (Ehrhardt et al., 2007). Recent evidence suggests influenza

A virus infection may up-regulate Akt activity during the early phase of infection, to

prevent rapid host cell death, then initiate apoptosis at later stages of infection (Zhirnov

and Klenk, 2007).

I.10.3 NS1 and virulence

The emergence of a highly pathogenic avian H5N1 influenza A virus in Hong

Kong has resulted in considerable activity to determine the viral factors that contribute to

its virulence. Using recombinant viruses, there are data that indicate that the viral proteins

HA, PB2 and NS1 can influence the degree of virulence of influenza A (Jackson et al.,

51

2008; Li et al., 2006). NS1 from highly virulent strains of influenza A has been shown to

be more effective at inhibiting a type I IFN response when compared to other less virulent

influenza A viruses. Virus strains without or with a defective NS1 cannot propagate in

cells with an intact type I IFN system, again highlighting the importance of IFNs in host

defence (Garcia-Sastre et al., 1998). Like other influenza virus proteins, there are many

variants of NS1 as a result of mutations, with several critical amino acid residues that

have been shown to correlate with virulence in the host. Glutamic acid substitution of

aspartic acid at position 92 in NS1 has been described in swine models to confer

resistance to various cytokines including TNF-α and IFN-α (Li and Wang, 2007; Seo et

al., 2004). Other studies indicate that an NS1 with a five amino acid deletion at positions

80-84, confers enhanced virulence to influenza A viruses, though the molecular basis of

this remains to be determined (Long et al., 2008).

52

I.11 Thesis hypothesis and objectives

Hypothesis: Influenza virus H5N1 NS1 interferes with IFN-α/ß inducible signal

transduction.

Objectives: To invetsigate the effects of NS1 on different aspects of type I IFN signaling

and evaluate the therapeutic potential of type I IFNs against H5N1 influenza

A infection.

53

CHAPTER II

MATERIALS AND METHODS

II.1 Cells, viruses and reagents

The human cervical carcinoma cell line HeLa was obtained from ATCC (Manassas, VA).

Cells were cultured in Dulbecco’s modified Eagle’s medium (Invitrogen), supplemented

with 10% fetal calf serum (FCS), 100 U/ml penicillin, 100 mg/ml streptomycin

(Invitrogen). Plasmid pBudCE4.1-H5N1 NS1-HA (A/Duck/Hubei/L-1) were kindly

provided by Dr. Bing Sun from Shanghai Pasteur Institute. Fresh lung biopsies were

obtained from patients having surgical resection of lung tissue in Queen Mary Hospital,

all of whom gave informed consent under a study approved by The Hong Kong

University and Hospital Authority (Hong Kong West) Institutional Review Board. The

biopsies or tissue fragments were excess to the requirements of clinical diagnosis. Non-

tumor lung tissue from each donor was cut into multiple fragments (2-3mm). The tissues

were immediately placed into culture medium (F-12K nutrient mixture with L-glutamine,

and antibiotics) and infected with influenza A H5N1 (A/Vietnam/3046/04) viruses within

three hours of collection. Influenza virus was used at a titer of 1×106 50% tissue culture

infectious doses (TCID50)/ml. Biopsies with no viruses were used as controls. The biopsy

or tissue fragments were incubated at 37 oC for 18 h. The DNase-treated mRNA from

infected ex vivo organ culture was extracted using RNeasy mini kit (Qiagen Hilden,

Germany). The cDNA as synthesized from cDNA using oligo-dT primers and Superscript

III reverse transcriptase (invitrogen). The infection of primary human lung tissue with

influenza A virus were performed by Dr. John Nicolls laboratory (University of

54

HongKong, China). Human recombinant IFN alfacon-1 (IFN alfacon-1, specific activity,

6×108 U/mL) was provided by Intermune (Brisbane, CA). Human IFN-β (specific

activity, 1.2×107 U/mL) was generously provided by Darren Baker (Biogen Inc., MA).

Antibodies against p-STAT1, p-STAT2, p-STAT3, STAT1, STAT3, HA, tubulin, SOCS1

and SOCS3 were purchased from Cell signaling. Antibodies against STAT2, Histone H1

and β-actin were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Secondary

HRP-conjugated goat anti-mouse antibody and HRP-conjugated goat anti-rabbit antibody

were purchased from GE healthcare Limited (UK).

II.2 Transfection and virus infection

Cells (2×105) were transfected using Lipofectamine LTX (Invitrogen, CA)

according to manufacturer protocol. Briefly, cells were seeded in 6 well plates 24 hours

before transfection. Plasmid DNA and transfection reagent were mixed in serum-free

media and incubated for 30 minutes at room temperature. Tranfection complexes were

then added gently to the 6 well plate. For H5N1 influenza A(A/Vietnam/3046/04)

infection, primary lung tissues were first pre-treated with either PBS or IFN alfacon-

1(1×104

U/mL) for 24 hours, tissue were then incubated with virus (1×106 TCID/mL) in

24 well plate for 30 minutes, tissues were subsequently washed twice with PBS and

placed in F12K medium for 18 hours. The infection of primary human lung tissue with

influenza A virus were performed by Dr. John Nicolls laboratory (University of

HongKong, China)

55

II.2 Immunoblotting and immunoprecipitation

Cells were lysed in 50 µl of lysis buffer (1% Triton X-100, 0.5% Nonidet P-40,

150 mM NaCl, 10 mM Tris-HCl, pH 7.4, 1 mM EDTA, 1 mM EGTA, 0.2 mM

phenylmethylsulfonyl fluoride). Protein concentration in lysates was determined using the

Bio-Rad protein DC assay kit (BioRad laboratories, Hercules, CA). 40 µg of protein

lysate/sample was denatured in 5x sample buffer and resolved by sodium dodecyl sulfate–

polyacrylamide gel electrophoresis (SDS-PAGE). The

separated proteins were transferred

to a nitrocellulose membrane, followed by blocking with 5% bovine serum albumin (w/v)

in TBS-T for 1 h at room temperature. Membranes were probed with the

specified

antibodies overnight. Membrane was the washed three times (5 minute/each time) with

1X TBS-T buffer. Membrane was then probed with appropriate secondary antibodies.

Proteins were visualized using the ECL detection system (Pierce, Rockford, IL). For

immunoprecipitations, cells transfected with vector alone or H5N1 NS1 plasmid were

incubated in hypotonic lysis buffer containing 10 mM HEPES, pH 7.4 for 30 minutes on

ice and the suspension was then briefly sonicated. The suspension was incubated for 30

minutes at 4°C, followed by centrifugation at 14 000 rpm for 30 min at 4°C. The

supernatant was collected and protein concentration was measured using the Bio-Rad

protein assay kit. 500µg of proteins were incubated with 1µg of either anti-STAT1

antibody or anti-STAT2 antibody or IgG isotype control antibody. Immune complexes

were immunoprecipitated with protein A/G-sepharose beads (Santa Cruz Biotechnology)

and washed 6 times with HEPES buffer. Beads were then denatured in 5x sample

reducing buffer and resolved by SDS-PAGE.

56

II.3 Cell sorting and flow cytometric analysis

24 hours post transfection, cells were first washed with PBS and subsequently

incubated with versene for 15 minutes. Suspended cells were collected and centrifuged at

1500 rpm. Cell pellets were then resuspended at 1×106 cells per 100µL of 2% FCS/PBS.

5×105 GFP

+ sorted cells were incubated with either anti-human IFNAR1 (Biogen) or anti-

human IFNAR2 antibody (Caltag), followed by PE-conjugated anti-mouse IgG antibody.

As isotype controls, cells were incubated with PE-labeled isotype control IgG antibody

(eBioscience). All analyses were performed using the FACSCalibur and CellQuest

software. Cells were gated based on forward and side scatter.

II.4 RNA extraction and cDNA synthesis

Cells were lysed using buffer RLT+β mercaptoenthanol with Qiagen QIA-

shreddar columns. RNA isolations were carried out using Qiagen RNeasy mini kit

according to the manufacturer’s protocol, including on column DNA digestion. Total

cellular RNA was eluted in RNase-free water. The concentration of RNA was

subsequently determined by UV spectrophotometry at 260nm wavelength (Beckman).

cDNAs were synthesized using 0.5µg RNA according to the manufacturer’s protocol

(Invitrogen). cDNA from primary human lung tissue were provided by Dr. John Nicolls

laboratory (University of HongKong, China)

II.5 Real time-polymerase chain reaction (RT-PCR)

Real-time PCR were performed using a LightCycler (Roche) in conjunction

with LightCycler FastStart DNA Master SYBR Green PLUS I Kits (Roche). Reactions

57

were performed in 20 µL containing 4µL Master SYBR GreenPLUS buffer at a final

concentration of 1X, 5µL of 0.1µg/µL cDNA. 9µL of PCR grade water and 1µL of each

20 µM forward and reverse primers. PCR reactions were performed under the following

conditions: pre-incubation at 95°C for 10 minutes, followed by 45 amplification cycles of

denaturation for 10 seconds, annealing for 5 seconds, extension at 720 for 10 seconds,

melting curve analysis at 650 for 15 seconds and a continuous acquisition mode of 95

0

with temperature transition rate of 0.1 per second. The data were subsequently analysed

using software RealQuant. Primers for IFN-α2 (PPH00379A-200) and IFN-β

(PPH00384E-200) were purchased from SA Bioscience (Frederick, MD). Other PCRs

were performed using the following primers:

IFNAR1

(forward) 5’ CACTGACTGTATATTGTGTGAAAGCCAGAG 3’

(reverse) 5’ CATCTATACTGGAAGAAGGTTTAAGTGATG 3’

IFNAR2

(forward) 5’ ATTTCCGGTCCATCTTATCAT 3’

(reverse) 5’ACTGAACAACGTTGTGTTCC 3’

Influenza A M gene

(forward) 5’ CTTCTAACCGAGGTCGAAACG 3’

(reverse) 5’ GGCATTTTGGACAAAGCGTCTA 3’

ISG15

(forward) 5’ TCCTGGTGAGGAATAACAAGGG 3’

(reverse) 5’ CTCAGCCAGAACAGGTCGTC 3’

58

PKR

(forward) 5’ GCCTTTTCATCCAAATGGAATTC 3’

(reverse) 5’ GAAATCTGTTCTGGGCTCATG 3’

2’5’OAS

(forward) 5’ AGCTTCATGGAGAGGGGCA 3’

(reverse) 5’ AGGCCTGGCTGAATTACCCAT 3’

ββββ-actin

(forward) 5’ACATGGAGAAAAATCTGGCAC 3’

(reverse) 5’ GTAGCACAGCTTCTCCTTAATGT 3’

II.6 Electrophoretic mobility shift assay

10µg of nuclear protein from untreated or IFN-treated cells were extracted as

described previously(Brierley et al., 2006). Extracts were incubated with 1µg poly(dI-

dC)poly(dI-dC) for ten minutes at 4°C in buffer containing 60mM EGTA, and 5% Ficoll

(final volume 30µl). 40,000 counts per minute (cpm) of radiolabeled SIE (5’-

AGCTTCATTTCCCGTAAATCCCT) were added and the reaction mixture was

incubated for an additional 20 minutes at ambient temperature. Protein-DNA complexes

were resolved on a 4.5% polyacrylamide gel using 0.5X TBE (final concentration 45mM

Tris borate, 1mM EDTA) as running buffer. Gels were dried and exposed to

autoradiographic film (Kodak BioMax MS) overnight at -80°C.

59

II.7 Immunohistochemistry and Confocal microscopy

Cells transfected with H5N1 NS1 plasmid were stained as described previously

(Rahbar et al., 2006). Various proteins were visualized using fluorescence-conjugated

secondary antibodies (Alexafluor-488: green, Alexafluor-555: red and Alexafluor 647:

blue) (Amersham Biosciences,Cardiff, United Kingdom). Images were collected using an

upright Leica SP2 confocal laser-scanning microscope (Leica Microsystems Heidelberg

GmbH, Mannheim, Germany), a 100x oil immersion lens (1.4 numerical aperture), and a

x4 digital zoom. Laser excitations were 488 nm (Ar/Kr) and 543 nm (He/Ne), attenuated

to 10% and 50%, respectively, by way of an acoustic-optical transmission filter.

Sequential scan mode was used to eliminate cross talk of detected signals, which were

filtered between 500 to 530 nm and 560 to 660 nm. Image resolution was 512 dpi by 512

dpi (12 bit), and line averaging (4x) was used. Optical sections were collected at 0.5µm

intervals through the entire cell.

60

CHAPTER III

RESULTS

III.1 Influenza virus H5N1 NS1 localizes primarily in the nucleus of HeLa cells

In a first series of experiments, plasmid encoding HA-tagged H5N1 NS1 was

introduced by transfection into HeLa cells. Cells were fixed and stained at 8 and 24 hrs

post transfection, and analyzed for NS1 expression using a fluorescent conjugated anti-

HA antibody and immunofluorescence microscopy. Expression of H5N1 NS1 was

observed at 8 hrs post transfection, and the data reveal that NS1 localized predominantly

in the nucleus, detectable in the cytoplasm at 24 hrs (Figure III.1).

61

Figure III.1 H5N1 NS1localizes predominantly in the nucleus of HeLa cells.

HeLa cells were transfected with HA-tagged H5N1 NS1plasmid. HeLa cells

were fixed and stained for HA (Red) at different time points post transfection

and analyzed using immunofluorescence microscopy. Data are representative of

two independent experiments.

62

Time post transfection (hours)

0 8 24

63

III.2 H5N1 NS1 expression inhibits both IFN-inducible STAT phosphorylation and

nuclear translocation

To determine the effect of H5N1 NS1 protein expression on type I IFN signaling,

HeLa cells were transfected with either vector alone or plasmid containing the H5N1 NS1.

24 hours post transfection, cells were serum starved, then either left untreated or treated

with IFN-β (1×103 U/mL) for 15 minutes. Protein extracts were collected and the IFN-

inducible activation of STAT proteins was analyzed by immunoblotting. In contrast to

cells transfected with vector alone, there was a notable reduction in IFN-inducible

STAT1, STAT2 and STAT3 phosphorylation in cells expressing H5N1 NS1 (Figure

III.2A). To examine the effect of NS1 on IFN alfacon-1 inducible STAT phosphorylation,

nuclear and cytoplasmic fractions were isolated post IFN alfacon-1 (1×103 U/mL)

stimulation and analyzed by immunoblot. In comparison to vector transfected cells, the

amount of phosphorylated STAT proteins was reduced in the presence of H5N1 NS1

protein, though the extent of reduction was less when compared to IFN-β treatment

(Figure III.2B). Furthermore, to confirm that this inhibition of IFN-inducible STAT

phosphorylation occurred only in cells expressing NS1, in an identical series of

transfection experiments cells were fixed and stained with both anti-phospho-STAT2 and

anti-HA (NS1) antibodies. In contrast to cells that lack H5N1 NS1 expression, which that

exhibit strong IFN-inducible phospho-STAT2 staining in the nucleus, we identified a

notable reduction in IFN-inducible phospho-STAT2 staining in H5N1 NS1-expressing

cells following IFN-β stimulation (Figure III.2C).

64

Figure III.2 H5N1 NS1 expression inhibits IFN-inducible STAT

phosphorylation. A) HeLa cells transfected with vector alone (■) or HA-tagged

NS1 plasmid (■) were left untreated (-) or treated (+) with IFN-ß (1×103

U/mL)

for 15 minutes, 24 hours post transfection. Cells harvested, and lysates were

resolved by SDS-PAGE and immunoblotted with the indicated anti-phospho-

STAT1, anti-phospho-STAT2, anti-phospho-STAT3 and anti-HA(NS1)

antibodies. Membranes were also stripped and reprobed with anti-tubulin, anti-

STAT1, anti-STAT2 and anti-STAT3 as loading controls. Relative fold

induction of phosphorylated STAT proteins were calculated using signal

intensity of phosphor-STATs over total STATs and normalized with untreated,

vector transfected cells. Data are representative of three independent

experiments. B) HeLa cells transfected with vector alone (■) or HA-tagged NS1

plasmid (■) were left untreated (-) or treated (+) with IFN-ß (1×103

U/mL) for 15

minutes 24 hours post transfection. Cytoplasmic and nuclear fractions were

purified and resolved by SDS-PAGE, then immunoblotted with the indicated

anti-phospho-STAT1, anti-phospho-STAT2 and anti-phospho-STAT3

antibodies. Membrane were also probed with anti-HA antibody to confirm

expression of NS1 protein, and anti-β-actin and anti-histone H1 were used as

loading controls for cytoplasmic fraction and nuclear fraction, respectively.

Relative fold induction of phosphorylated STAT proteins was calculated using

signal intensity over loading and normalized with untreated, vector transfected

cells. Data are representative of two independent experiments. C) H5N1 NS1

expression inhibits IFN-inducible nuclear translocation of phosphorylated

STAT2. HeLa cells transfected with HA-tagged NS1 plasmid were treated with

IFN-ß (1×103 U/mL) for 15 minutes. Cells were then fixed and stained for HA

(red) and phospho-STAT2 (blue), and analyzed by confocal micrscopy as

described in Materials and Methods. Data are representative of two independent

experiments.

65

66

B

67

C

NS1 Merge p-STAT2

68

III.3 H5N1 NS1 protein expression inhibits IFN-inducible STAT:sis-Inducible

Element (SIE) complex formation

STAT proteins, once activated by phosphorylation, will form functional homo- or

hetero-dimeric complexes that migrate to the nucleus and target specific promoter

elements to initiate transcriptional activation of ISGs. To examine the functional

consequence of H5N1 NS1-mediated inhibition of IFN-inducible STAT phosphorylation,

electrophoretic mobility shift assay (EMSA) studies were carried out. Specifically, HeLa

cells transfected with empty vector or vector containing NS1, were either left untreated or

treated with IFN-β (1×103 U/mL) for 15 mins, 24 hours post transfection. Nuclear

extracts were prepared, incubated with radiolabeled sis-inducible elements (SIE), then

resolved by agarose gel electrophoresis to determine the formation of IFN-inducible

STAT:SIE complexes . In contrast to cells transfected with empty vector, we observed a

notable reduction in STAT3:3:SIE, STAT1:1:SIE and STAT1:3:SIE complexes in the

presence of H5N1 NS1 protein (Figure III.3).

69

Figure III.3 H5N1 NS1 expression inhibits IFN-inducible STAT:SIE

complexes formation. HeLa cells transfected with either vector alone or

HA-tagged NS1 plasmid were left untreated (-) or treated (+) with IFN-ß

(1×103U/mL) for 15 minutes 24 hours post transfection. Nuclear extracts

were isolated and incubated with 32

P-labeled SIE probe. Complexes were

resolved by native gel electrophoresis and visualized by autoradiography.

Data are representative of two independent experiments.

70

71

III.4 H5N1 NS1 expression leads to a reduction in IFNAR1 but not IFNAR2 cell

surface expression

The inhibition of IFN-inducible STAT phosphorylation prompted us to investigate

the influence of NS1 expression on upstream molecules involved in type I IFNs signaling,

starting with the receptors, IFNAR1 and IFNAR2. HeLa cells transfected with vector-

GFP or NS1-GFP plasmid were sorted 24 hours post transfection and analyzed for

surface IFNAR1 and IFNAR2 expression using anti-IFNAR1 and anti-IFNAR2

antibodies and flow cytometry. Surprisingly, cells expressing H5N1 NS1 exhibit reduced

level of surface IFNAR1 when compared to cells expressing vector alone (Figure III.4A).

Notably, IFNAR2 expression was not affected by the expression of NS1. In a subsequent

series of experiments we confirmed that this reduction in cell surface IFNAR1 expression

was restricted to cells expressing NS1 (Figure III.4B).

To determine whether the differential surface expression of IFNAR1 and IFNAR2

in the presence of H5N1 NS1 is a consequence of regulation at the mRNA level, RNA

was extracted from HeLa cells transfected with either vector-GFP or NS1 GFP plasmid

24 hours following transfection. IFNAR1 and IFNAR2 gene expression was analyzed

using real-time PCR. In contrast to vector-GFP transfected cells, we observed a reduction

in IFNAR1 but not IFNAR2 gene expression in cells transfected with H5N1 NS1 (Figure

III.5A).

To determine if this selective inhibition occurs in the context of H5N1 influenza

A infection , mRNA from primary human lung cells that were either mock infected (PBS)

or infected with H5N1 influenza A virus were collected and analyzed using real-time

PCR. Briefly, primary human non-tumor lung tissue obtained from patients undergoing

72

lung resection were isolated and infected with influenza A H5N1 virus, as described in

Materials & Methods. At 18hrs post-infection, RNA were extracted from the lung tissues

and used to generate cDNA. Infection with influenza A H5N1 virus resulted in a selective

reduction in IFNAR1 but not IFNAR2 gene expression when compared to controls

(Figure III.5B).

73

Figure III.4 Expression of H5N1 NS1 reduces surface IFNAR1 but not

IFNAR2 expression. A) HeLa cells were transfected with either GFP vector

alone (green) or GFP vector containing HA-tagged NS1 gene (red). 24 hours

post transfection, GFP+ cells and GFP

+NS1

+ cells were sorted by flow

cytometry 24 hours, and their surface IFNAR1 or IFNAR2 expression were

analyzed by flow cytometry. Data are representative of three independent

experiments. B) HeLa cells transfected with HA-tagged NS1 plasmid were

fixed and stained for HA (green) and either IFNAR1 (red) or IFNAR2 (red)

24 hours post transfection, then subsequently analyzed by confocal

microscopy. Data are representative of two independent experiments.

74

75

Figure III.5 A) Expression of H5N1 NS1 reduces IFNAR1 but not IFNAR2

mRNA expression. HeLa cells were transfected with either GFP vector alone

( ) or GFP vector containing HA-tagged NS1( ). GFP+ cells were sorted by

flow cytometry, RNA extracted and cDNA synthesized. Gene expression for

IFNAR1, IFNAR2 and ß-actin was measured by real-time PCR analysis.

Relative percentages of mRNA reduction were normalized using ß-actin gene

expression as loading control. Data are representative of two independent

experiments. B) RNA from primary human lung cells either mock infected with

PBS ( ) or infected with H5N1 influenza A virus ( ) was harvested 18 hours

post infection. Gene expression for IFNAR1, IFNAR2 and ß-actin was

measured by real-time PCR analysis. Relative gene expression was calculated

relative to ß-actin gene expression and normalized to mock infected cells. Data

are representative of two independent experiments.

Note: Infection of primary human lung cells, RNA extraction, cDNA synthesis

were performed by John Nicolls (University of HongKong)

76

77

III.6 IFNAR1 mRNA transcript has a longer half-life than IFNAR2 mRNA

The reduction in IFNAR1 but not IFNAR2 gene expression in the presence of

H5N1 NS1 suggests that there is a selective inhibition against IFNAR1, but it is also

possible that both IFNAR1 and IFNAR2 gene expression are suppressed in the presence

of H5N1 NS1, and the reduction in IFNAR1 that are detected were contributed by its

shorter mRNA half-life. To compare the intrinsic mRNA half-life between IFNAR1 and

IFNAR2, HeLa cells were treated with α-amanitin (2µg/mL), an inhibitor of RNA

polymerase II & III in a time course study. RNAs were collected at different time point

post treatment and used for cDNA synthesise. PCR analysis of IFNAR1 and IFNAR2

gene expression revealed that the turnover rate of IFNAR1 transcripts was slower when

compared to that of IFNAR2 transcripts, indicating that the NS1-mediated reduction of

gene expression is selectively targeting IFNAR1 but not IFNAR2 (Figure III.6).

78

Figure III.6 IFNAR1 mRNA does not have a shorter half-life than IFNAR2

mRNA. Hela cells were incubated with α-amanitin (2µg/mL) for various

time points. Total RNA was isolated and used for cDNA synthesis. Gene

expression of IFNAR1 (■), IFNAR2 (■) and 18S rRNA was analyzed by

PCR. Relative percentages of mRNA expression was calculated relative to

18S rRNA and normalized to untreated controls. Data are representative of

two independent experiments.

79

0

20

40

60

80

100

120

0 5 10 15 20 25 30

IFNAR1

IFNAR2

18S rRNA

0

20

40

60

80

100

120

0

20

40

60

80

100

120

Rel

ativ

e gene

expre

ssio

ns

(%)

4 8 12 16 240 28

Time (hour)

4 8 12 240 Time (hour)α-amanitin

80

III.7 H5N1 NS1 expression induces up-regulation of SOCS1 but not SOCS3

To determine the effect of H5N1 NS1 protein expression on negative regulators of

type I IFN signaling, namely SOCS1 and SOCS3, HeLa cells transfected with either

vector alone or plasmid containing the H5N1 NS1 gene were lysed 24 hours post

transfection. Protein extracts were collected, and SOCS1 and SOCS3 expression were

analyzed by Western immunoblotting. In contrast to cells transfected with vector alone,

we observed an increase in SOCS1 but not SOCS3 expression in cells expressing H5N1

NS1 (Figure III.7).

81

Figure III.7 Expression of H5N1 NS1 increases SOCS1 but not SOCS3

expression. HeLa cells transfected with vector alone or HA-tagged NS1 plasmid.

24 hours post transfection, cells were harvested and lysates were resolved by SDS-

PAGE, then immunoblotted with the indicated anti-SOCS1 and anti-SOCS3

antibodies. Membranes were probed with anti-HA antibody to confirm expression

of NS1, and anti-tubulin for loading. Relative fold induction of proteins were

calculated using signal intensity over loading and normalized with vector

transfected cells (SOCS1 ■ , SOCS3 ■ ). Data are representative of two

independent experiments.

82

p > 0.05

83

III.8 IFN pretreatment up-regulates ISGS and inhibits H5N1 influenza A infection

in primary human lung cells

In a final series of experiments, using non-tumor human lung tissue, we examined

the effect of IFN-α, namely IFN alfacon-1, on H5N1 influenza A infection. Primary

human lung tissues were collected from patients undergoing non-tumor lung biopsy at

Queen Mary’s Hospital, HongKong. Tissues were either pretreated with media or IFN-

alfacon-1 for 30 minutes prior to H5N1 influenza A infection. 18 hours post infection,

RNA was extracted for cDNA synthesis. RT-PCR analysis of influenza A M gene

expression suggests IFN-alfacon-1 pre-treatment can effectively inhibit H5N1 influenza

A replication (Figure III.8A). Our analysis of various ISGs in human lung cDNA

revealed that infection with H5N1 failed to invoke notable transcriptional activation of

IFN-α and IFN-β (Figure III.8B). Gene expression analysis of 2’5’-OAS, PKR and

ISG15, ISGs associated with an IFN-inducible antiviral response, revelaed that the gene

expression levels for these ISGs were not significantly altered, consistent with the failure

of H5N1 infection to induce an IFN response. However, analysis of IFN-alfacon-1 treated

primary human lung tissues revealed that an upregulation of expression of ISGs in both

mock and H5N1 influenza A infected samples (Figure III.8B).

84

Figure III.8. IFN treatment inhibits H5N1 (A/Vietnam/3046/04) replication in

primary human lung cells. Primary human lung cells were either left untreated or

treated with IFN alfacon-1, IFN-β, or IFN-λ1 (10000 U/mL) for 30 minutes.

Tissues were then infected with H5N1 influenza A virus. 18 hours post infection,

RNA from cells was collected and cDNA synthesized. Expression of A) influenza

M gene, B) PKR ( ), ISG15 ( ), 2’5’-OAS ( ), IFN-α ( ), IFN-β ( ) and ß-

actin gene expression was measured by real-time PCR analysis. Data are

representative of two independent experiments. Fold induction of gene expression

was calculated relative to ß-actin gene expression and normalized to mock

infected controls.

Note: infection of primary human lung cells, RNA extraction, cDNA synthesis

and M gene expression analysis were performed by John Nicolls (University of

HongKong)

85

86

CHAPTER IV

DISCUSSION

The coordinated host response to virus infection involves activation of both innate

and adaptive immune responses. Notably, a robust innate immune response is critical and,

as described earlier, major effectors in the earliest innate immune response to infection by

virus are the type I IFNs, IFN-αs/β. Influenza A viruses have evolved to target type I

IFNs, largely facilitated by the NS1 protein (Kochs et al., 2007). Indeed, it is likely that

the IFN system is the primary target of NS1. Studies involving the use of influenza virus

mutants, specifically defective NS1 mutants, demonstrated that in the absence of a

functional NS1, influenza infection is significantly attenuated in cells with an intact IFN

system. Conversely, replication of an influenza virus with a non-functional NS1 can be

restored in cells or in mice if the IFN system is defective (Garcia-Sastre et al., 1998).

NS1 will inhibit dsRNA-mediated IRF3 phosphorylation and its translocation into

the nucleus, thereby blocking the transcriptional activation of IFNs-α/β (Talon et al.,

2000; Wang et al., 2000). However, the interaction between NS1 and the cytosolic sensor

RIG-I is primarily responsible for the blockade of type I IFN induction during influenza

infection (Guo et al., 2007; Opitz et al., 2007). Evidence from several groups indicates

that RIG-I is critical in regulating type I IFN expression in response to ssRNA bearing 5’

phosphates (Hornung et al., 2006; Pichlmair et al., 2006). Knockdown of RIG-I limits

IFN production during influenza infection, whereas overexpression of RIG-I can override

the inhibitory function of NS1, and block viral replication.

Herein we report a novel mechanism by which the H5N1 influenza virus NS1

alters IFN-α/β signalling, possibly mediated by the selective inhibition of IFNAR1 gene

87

expression. The inhibitory effects of NS1 on IFNs-α/β have been largely attributed to its

ability to inhibit IFN induction. Prompted by the increase of drug-resistant influenza A

isolates, we initiated experiments to determine the effect of NS1 on type I IFN signaling

as a way of exploring the therapeutic potential of type I IFNs during influenza A infection.

Expression of the H5N1 influenza virus NS1 in HeLa cells led to the reduction of IFN-

inducible STAT phosphorylation (Figure III.2). The phosphorylation-activation of

STAT1 and STAT2 is critical for mediating IFN-α/β responses (Fu et al., 1990; Levy et

al., 1989). In the absence of these transcriptional effector proteins, cells are unresponsive

to IFNs-α/β and are highly susceptible to virus infection (Durbin et al., 1996; Park et al.,

2000). IFN-inducible activation of the Jaks associated with IFNAR leads to the

recruitment and tyrosine phosphorylation-activation of STATs, then their subsequent

dimerization and translocation into the nucleus to activate gene expression via binding to

specific elements in the promoter regions of ISGs. Examination of IFN-inducible

STAT:SIE complexes, using EMSA, demonstrated a reduction in the formation of

STAT1:1 and STAT1:3 complexes in the presence of the influenza virus H5N1 NS1

(Figure III.3).

A number of viruses have evolved to target STAT proteins to block the antiviral

activity of IFNs. Paramyxoviruses such as SV5 and type II human parainfluenza viruses

(HPIV2) block IFN signalling, mediated by their V proteins, which induce proteasomal

degradation of STAT1 and STAT2 through polyubiquitation(Andrejeva et al., 2002;

Precious et al., 2005). HCV core proteins block STAT1 activation and subsequent

function, mediated by STAT1-core protein interactions and suppression of STAT1 gene

expression (Lin et al., 2006).

88

Having demonstrated that the expression of the influenza virus H5N1 NS1 protein

reduced the extent of IFN-inducible STAT phosphorylation and STAT-DNA binding, we

undertook experiments to determine whether NS1 directly associates with STAT proteins.

Immunoprecipitation studies did not reveal notable physical association between NS1 and

STAT proteins, suggesting the inhibition of IFN-inducible STAT signaling was likely not

a consequence of strong physical interaction between NS1 and STAT proteins. In

subsequent experiments, we undertook to evaluate whether upstream effectors of the

STAT proteins might be affected by NS1 expression. Flow cytometric analysis of

surface IFNAR1 and IFNAR2 expression revealed a reduction in cell surface IFNAR1 in

the presence of H5N1 NS1, but more interestingly, IFNAR2 surface expression remained

unaltered. Immunofluorescence confocal microscopy were consistent with these

observations (Figure III.4B). Based on the ability of NS1 to inhibit the generation of

mature host mRNA, experiments were conducted that revealed that the reduction in

surface expression of IFNAR1 was a consequence of a decrease in gene expression

(Figure III.5). From these results we infer that the NS1-dependent reduction in IFNAR1

gene expression likely results in a decrease in IFN-inducible STAT phosphorylation and

DNA binding. Indeed, in other studies with cardiac fibroblasts and myocytes, the higher

basal levels of IFNAR1, JAK1, TYK2, IRF9 and STAT2 in the cardiac fibroblasts

compared with the myocytes, correlated directly with a stronger IFN response (Zurney et

al., 2007). The importance of IFNAR1 in an IFN response is further supported by earlier

studies that showed that IFNAR1 null cells are non-responsive to IFN, and IFNAR1 null

mice are highly susceptible to virus infections (Hwang et al., 1995).Interestingly, in

clinical studies of HCV patients who did not respond or were less sensitive to IFN

89

therapy, evidence was provided for reduced levels of either IFNAR1 or IFNAR2 gene

expression compared to IFN responders (Morita et al., 1998). Polymorphisms in the

promoter region of IFNAR1 and IFNAR2 have been closely linked with susceptibility to

a number of diseases including malaria, multiple sclerosis, trypanosomaiasis, HCV and

HIV (Aucan et al., 2003; Diop et al., 2006; Leyva et al., 2005; Tena-Tomas et al., 2007).

In contrast, overexpression of IFNAR1 and IFNAR2, as is the case in Down’s Syndrome

patients, where chromosome 21 is trisomic, results in enhanced sensitivity to IFN

(Mowshowitz et al., 1983). Viewed altogether, the inhibitory effect of influenza virus

H5N1 NS1 on IFNAR1 gene expression is an effective mechanism to render target cells

less sensitive to IFN.

In the context of viral infections, IFNs-α/β not only act as critical components of the

innate immune response, but also play a prominent role in modulating the adaptive

immune response. Consequently, NS1-mediated inhibition of IFN signalling will have a

negative impact on the adaptive immune response. Viral clearance of influenza infection

is predominantly mediated by virus-specific CD8+ cytotoxic T cells (Doherty et al., 1997).

This process requires T helper 1 (Th1) cell activity, dependent on interactions with

antigen presenting cells (APCs) such as dendritic cells (DC). IFNs-α/β promotes the

differentiation and maturation of DCs (Santini et al., 2002; Santini et al., 2000).

Incubation of blood-derived monocytes with IFN-α induces maturation/activation

markers in DCs, e.g. CD83 and CD25, along with increased expression of the major

histocompatibility complex (MHC), costimulatory molecules (CD80 and CD86) and

chemokine receptors. Viral PAMP recognition by DCs leads to a series of downstream

responses including type I IFN production that contributes to DC maturation. Mature DCs

90

present viral peptide-MHC complexes along with other costimulatory molecules to

stimulate or prime virus-specific CD4+ and CD8+ T cells. IFNs-α/β also regulate the

production of several chemokines in DCs that have a role in the recruitment of NK cells,

activated or memory T cells, and pDCs to sites of infection (Lande et al., 2003; Padovan

et al., 2002). Defects in any aspect of IFN signaling can lead to defective DC maturation

and effector functions. Influenza A virus infection of DCs results in a block in DC

maturation and subsequent ineffective T cell activation, mediated by NS1 (Fernandez-

Sesma et al., 2006). NS1 expresssion in DCs altered the expression of a panel of genes

that were required for both maturation and migration, including IFN-α and IFNAR1

(Fernandez-Sesma et al., 2006). Consistent with this finding, we observe a reduction in

IFNAR1 gene expression in influenza virus H5N1 infected primary human lung cells. A

similar reduction in IFNAR1 gene expression was identified in HeLa cells expressing the

H5N1 NS1, suggesting the viral-induced downregulation of IFNAR1 gene expression in

the infected primary lung cells was NS1-dependent (Figure III.5). Based on current

knowledge of NS1, the reduction in IFNAR1 gene expression is likely mediated by

inhibition of pre-mRNA splicing and/or polyadenylation. NS1 interacts with components

of the splicing machinery, U6 snRNA. U6 snRNA has been shown to complex with other

constituents of the spliceosome in an orderly fashion to mediate pre-mRNA splicing (Qiu

et al., 1995). An association between NS1 and U6 snRNA hinders its ability to complex

with other catalytic subunits of the spliceosome, thereby leading to the accumulation of

pre-mRNAs in the nucleus of the host cell. Additionally, NS1 affects polyadenylation of

host mRNA through targeting CPSF30 and PABII (Chen et al., 1999; Nemeroff et al.,

1998). 3’ cleavage and polyadenylation of mRNAs promotes their export into the

91

cytoplasm, whereas mRNAs that have undergone 3’ cleavage alone are retained in the

nucleus (Fuke and Ohno, 2008).

It is noteworthy that, using a distinct strategy to the NS1 protein, vaccinia virus

encodes a type I IFN receptor homologue that functions to prevent IFN signaling by

functioning as a decoy receptor and sequestering IFN away from cell surface receptors

(Colamonici et al., 1995).

Notably, expression of H5N1 NS1 was shown to target the expression of IFNAR1

but not IFNAR2 (Figure III.5). We provide further evidence to suggest that this selective

inhibition is not an indirect effect of their differential mRNA half-life (Figure III.6). The

mechanism behind this selective inhibition in gene expression is currently under

investigation. Though promoter analysis of IFNAR1 and IFNAR2 suggests they contain

elements that can respond to type I IFNs, regulation of IFNAR1 and IFNAR2 gene

expression has not been closely examined. IFNAR1 gene expression has been

demonstrated to be lower in patients suffering from multiple sclerosis, and long term IFN

therapy seems to increase IFNAR1 gene expression (Serana et al., 2008). In addition,

HCV patients that are undergoing IFN therapy exhibit an increased gene expression for

IFNAR2 when compared to naïve controls (Fujiwara et al., 2004).

Interestingly, several studies that have employed gene microarray analysis of

intact or mutant NS1 transfected cells provide evidence to suggest that only a selective

pool of genes are affected in the presence of intact NS1 (Fernandez-Sesma et al., 2006).

Though NS1 disrupts numerous post-transcriptional modifications, reduction in gene

expression does not seem to be a global effect, and the expression of many genes remain

92

up-regulated or unaffected in the presence of NS1 (Fernandez-Sesma et al., 2006;

Shimizu and Kuroda, 2004, 2006).

Furthermore, given IFNs signaling can be also be negatively regulated by both

SOCS1 and SOCS3 expression, we undertook a study to examine whether their

expression levels are affected by the H5N1 NS1 (Liu et al., 2004; Song and Shuai, 1998).

We provide evidence that H5N1 NS1 expression can upregulate SOCS1, but not SOCS3,

24 hours post transfection (Figure III.7) Recent evidence from studies with H1N1

suggests that influenza infection can inhibit type I IFN signaling through up-regulation

of SOCS3 expression, yet, there was no evidence for the H1N1 NS1 mediating this effect.

These apparent contradictory data with the H1N1 infection data may be reflective of

structural differences between the two NS1 proteins: whereas the H1N1 NS1 protein

dimerizes in solution, H5N1 NS1 has distinct structural features and can form oligomers

in solution (Bornholdt and Prasad, 2008).

Despite the strong inhibitory mechanism employed by H5N1, we show that

exogenous IFN treatment can over-ride these effects, and effectively up-regulate a

number of ISGs to establish an anti-viral state within cells (Figure III.8). Current clinical

therapeutic strategies for patients with influenza A infections include the inhibitors of the

Adamantine family, which inhibit the M2 ion channel to block the release of vRNP into

the cytoplasm (Davies et al., 1964; Wang et al., 1993), and those that target the NA

protein to prevent viral budding (De Clercq, 2004; Moscona, 2005). Despite their earlier

successes, the frequency of influenza A virus isolates exhibiting resistance to these

inhibitors has increased dramatically, prompting a need for alternative antiviral

therapeutic strategies (Bright et al., 2005; Bright et al., 2006). Herein, we demonstrate

93

that IFN alfacon-1 can effectively block H5N1 influenza A replication in human lung

cells, warranting further investigations into the effective use of IFNs as a viable

therapeutic strategy to control H5N1 influenza A infection in humans.

The expression of IFNAR1 and IFNAR2 has not been scrutinized in the context of

other strains of influenza A or NS1-expressing cells, thus it is difficult to surmise whether

such selective inhibition of IFNAR1 is exclusively associated with the highly pathogenic

strain like H5N1, or a conserved phenomenon across strains of influenza viruses.

However, studies using recombinant virus strongly suggest NS1 plays a prominent role in

dictating the virulence of influenza viruses. Substitution of NS1 derived from highly

pathogenic strains such that of H5N1 or the 1918 H1N1 into the low virulence influenza

viruses greatly enhances their pathogenicity in animals (Jackson et al., 2008; Li et al.,

2006). Mutagenesis experiments suggest a number of key residues in NS1 are closely

associated with virulence. Substitution of aspartic acid to glutamic acid at position 92 has

been shown to confer resistance to antiviral cytokines like TNF-α and IFN-α in swine

models of infection (Jackson et al., 2008; Li et al., 2006). Pre-treatment of cells with

these antiviral cytokines did not block subsequent viral replication, and the underlying

mechanism of action remains unknown. Another cluster of amino acids in NS1 that

apparently contribute to virulence are five amino acids at positions 80-84. Deletion of

these amino acids in NS1 has been shown to increase NS1’s effectiveness as an IFN

antagonist. The H5N1 NS1 in our study was negative for the D92E mutation but it did

possess the 80-84 amino acid deletion, from which we might infer that this deletion may

potentially influence the reduction in IFNAR1 gene expression.

94

As the number of drug-resistant influenza isolates increases, much attention is

being focused on NS1, in the hope of developing novel therapeutics against influenza

infection (Sheu et al., 2008). Certainly, insights into the mechanism of action of NS1 as a

virulence factor, as described herein, confirm that disabling NS1 is a reasonable strategy

for the development of antiviral drugs targeted against influenza A viruses.

95

CHAPTER V

FUTURE DIRECTIONS

The inhibition of IFN-inducible STAT activation in the presence of H5N1 NS1

was examined only in the context of tyrosine phosphorylation. However, complete

transcriptional activation of STAT proteins requires both phosphorylation on tyrosine as

well as serine residues. Thus it would be of interest to examine the effect of H5N1 NS1

on IFN-inducible serine phosphorylation.

Another question not addressed in our studies relates to the effect of H5N1 on the

gene expression ratio between different IFNAR2 isoforms. IFNAR2 has three isoforms

derived from pre-mRNA splicing, with only IFNAR2c being the functional receptor that

can transduce signal upon ligand binding. Ratios between functional and non-functional

receptors can influence the strength and kinetics of IFN signaling. Because the flow

cytometric analysis, immunofluorescence and RT-PCR studies did not allow us to

distinguish among the different isoforms, subsequent experiments to examine the ratio of

the different IFANR2 isoforms in the presence of H5N1 NS1 would provide insights into

the effects of NS1 on IFNAR2 gene expression.

NS1 mutation experiments are planned to identify the domain(s) or amino acid

residues responsible for the reduction in IFNAR1 gene expression. Several key residues

have been shown to play important roles in the activity of NS1 in suppressing IFN

responses, including Arg 38 and Ser 42 in the dsRNA binding domain, and Phe 103 and

Met 106 in the C-terminal domain (Kochs et al., 2007). Using site-directed mutagenesis,

residues 38, 42, 103 and 106 will be targeted, then the effects of expression of these

96

mutant NS1 constructs in HeLa cells examined in the context of IFNAR1 gene and cell

surface expression.

97

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