INVESTIGATING RNA GRANULES

FORMATION DURING

CALICIVIRUSES INFECTION

By

MAJID NOORI HUMOUD AL-SAILAWI

THESIS SUBMITTED FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

FACULTY OF HEALTH AND MEDICAL

SCIENCES

UNIVERSITY OF SURREY

May 2015

i

ABSTRACT

Human norovirus (HuNV) is a member of the calicivirus family and is a major cause of

viral gastroenteritis worldwide. Due to the absence of a suitable cell culture system, HuNoV

replication mechanisms are poorly understood, but two animal caliciviruses, Feline

calicivirus (FCV) and Murine Norovirus (MNV) provide models to increase our

understanding of norovirus biology. Unlike cellular mRNAs, the calicivirus RNA genome

does not possess a 5' cap structure but instead has a 13–15 kDa viral protein, genome linked

(VPg) directing translation, hijacking the host protein synthesis machinery. The viral life

cycle requires separated events occurring at different times since viral transcripts are used as

the template both for translation (mRNA) and replication (genomic RNA). Therefore

mechanisms are required to control the viral RNA fate. In eukaryotes, during stress

conditions, mRNAs can be stored in subcellular compartments such as stress granules to stall

their translation or in processing bodies to be degraded. Recent evidence indicates that these

compartments also play an important role during the viral life cycle. Therefore, using

immunofluorescence microscopy we set out to investigate how FCV and MNV infection

regulate the formation of G3BP1- and PABP-1-containing stress granules and DCP-1-

containing processing bodies to address whether these cytoplasmic granules could play a role

during the viral life cycle. We have now shown that FCV has the ability to prevent stress

granules formation during infection and that this is important for replication in CRFK and

FEA cells. Using FCV-free supernatant from infected CRFK cells and immunofluorescence

microscopy, we have also shown that during infection, the formation of stress granules is

induced in a paracrine manner in uninfected cells via a messenger molecule released from

infected cells. We hypothesize that this could reflect a new antiviral role for stress granules.

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Furthermore, MNV and FCV infection also led to the disruption of processing-bodies

assembly. Overall, this study revealed that caliciviruses modulate the RNA granules during

infection and that this could be part of viral mechanism to counteract the antiviral response.

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ACKNOWLEDGEMENTS

I would like to say a huge thank you to my supervisor Dr Nicolas Locker for his

advice, endless patience, guidance, inspiration and support. It has been pleasure to working

under his supervision and I am lucky to have worked with him. Without his supervision and

constant help my dissertation would not have been possible.

I would also like to thank my co-supervisor Professor Lisa Roberts for her guidance,

encouragement and support. I am extremely grateful to her.

A massive thank you also goes to Dr Elizabeth Royall for her assistance, support, proof

reading and brilliant comments and suggestions. I also need to thank Dr Nicole Doyle and Dr

Margaret Carter. Thank you for all your help, support and your friendship. I would also like

to thank Dr Dominique Weill in Université Paris 6 (Paris, France) for her advice. I would also

like to thank Dr. Belinda Hall for her advice. Thanks also to Dr Rachel Butler and Robert

Francis for their help and advice with the confocal work.

To my lab-mates over the years, Dr Melvin Leteane, Dr Azimah, David, Salmah, Nargs,

Mariam, Nefeli and Hanan, a big thank you for the all fun times, your friendship and support.

Thank you to the University of Basra for proving funding for my study.

My heartfelt thanks to my extended family and my wife family for their encouragement,

prayers and helping me over these four years.

To soul of my father and my granny, thank you to painting me in the right way. I miss you.

Finally to my mother, brother and his family, my sister, my wife and my children Mohammed

Ridha and Yusuf. You have lived through all the ups and downs of this PhD with me, and for

have given me your constant love, support and encouragement. I could not have done this

without you (no thank you can ever be enough).

It’s all for you.

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TABLE OF CONTENT

CHAPTER 1 Introduction 1

1.1 RNA Granules 2

1.2 Cytoplasmic mRNPs granules (Stress granules and P-bodies) 3

1.3 Stress granules (SGs) 4

1.3.1 Stress granule composition 6

1.3.2 Stress granule assembly/disassembly 11

1.3.3 Functions and importance of stress granules 17

1.4 Processing bodies (P-bodies or PBs) 20

1.4.1 P-body assembly/disassembly 23

1.4.2 Processing bodies functions 25

1.4.3 Role of PBs in mRNA degradation 26

1.4.4 Role of P-bodies in mRNA surveillance 28

1.4.5 Role of P-bodies in gene silencing 28

1.4.6 Other functions of PBs 29

1.5 Interactions between Stress granules and P-bodies 30

1.6 Stress granules, P-bodies, and viral life cycles 35

1.6.1 Viral interaction with stress granules 36

1.6.2 Inhibition of Stress granules by viral proteinase 38

1.6.3 Inhibition of stress granule formation by modulation of eIF2α phosphorylation 39

1.6.4 Inhibition of stress granule formation by sequestering or co-opting their

components

40

1.6.5 Viral replication mechanism that benefits from SG formation 43

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1.7 Virus/P-bodies interaction 45

1.8 Caliciviruses 48

1.8.1 Lagoviruses (LaVs) 51

1.8.2 Vesiviruses (VeVs) 52

1.8.3 Sapoviruses (SaVs) 53

1.8.4 Neboviruses (NeVs) 55

1.9 Noroviruses (NoVs) 55

1.9.1 Animal noroviruses 57

1.9.2 Murine norovirus-1 (MNV-1) 57

1.9.3 Human noroviruses (HuNoVs) 58

1.9.4 Clinical signs 61

1.9.5 Transmission 62

1.10 Model systems for the study of human norovirus biology 63

1.11 Pathogenesis and immunity 67

1.12 Viral structure, genome organisation and function of the viral proteins 70

1.13 Norovirus life cycle 78

1.14 The function of RNA-protein interaction during noroviruses infections 84

1.15 Aims and objectives of the research 88

CHAPTER 2 General Materials and Methods 89

2.1 Maintenance of cells 90

2.1.1 Crandall Rees Feline Kidney (CRFK) Cells 90

2.1.2 Mouse Leukemic monocyte-macrophage Cells (RAW 264.7) 90

2.1.3 Feline embryonic airway (FEA) cells 91

2.1.4 Murine microglial cells (BV-2) 91

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2.1.5 Human embryonic kidney 293 (HEK293T) cells (293 cells) 92

2.1.6 Murine macrophage J774 cells 92

2.1.7 THP-1 cells 93

2.2 Freezing and Resuscitation of CRFK and RAW264.7 cells 93

2.3 Preparation of viruses stock 94

2.3.1 Preparation of Feline Calicivirus (FCV) stock 94

2.3.2 Production of Murine Norovirus (MNV-1) stock 94

2.4 50% Tissue Culture Infectious Dose (TCID50) Assay 95

2.5 Preparation of VPg-linked FCV RNA 95

2.6 Preparation of VPg-linked MNV RNA 96

2.7 Preparation and quantitation of VPg-linked FCV and MNV RNA 97

2.8 Immunofluorescence studies 97

2.8.1 Cell culture and virus infection 97

2.8.2 Immunofluorescence assay 97

2.9 Effect of calicivirus infection on SGs and PBs formation 98

2.9.1 Staining for FCV-p76 protein 100

2.9.2 Staining for MNV-3D polymerase protein 100

2.9.3 Staining for SG-markers 100

2.9.4 Staining for PBs-markers 101

2.10 SGs formation during FCV infection 101

2.10.1 Treatment with sodium arsenite (NaAsO2) 101

2.10.2 Effect of FCV infection on SGs formation 102

2.10.3 Effect of NaAsO2–induced SGs on FCV replication 102

2.10.4 Effect of FCV infection on hydrogen peroxide (H2O2) induced SGs 102

2.10.5 Effect of the H2O2-induced SGs on FCV replication 103

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2.11 Preparation of FCV free supernatants 103

2.11.1 Virus infection and clarification (virus elimination) 103

2.11.2 Virus Precipitation 105

2.13 Virus inactivation using ultraviolet-light (UV-light) 105

2.14 Effect of cell culture supernatant on stress granules formation 106

2.15 Effect of interferon on stress granules formation 106

2.16 Effect of RNase A treatment on stress granules formation 106

2.16 Effect of heat shock on stress granules formation 107

2.17 SGs formation during MNV-1 infection 107

2.18 Formation of P-bodies during FCV Infection. 108

2.19 Formation of P-bodies during MNV-1 Infection 108

2.20 Statistical analysis

108

CHAPTER 3 Regulation of Stress Granules formation during FCV and MNV-1

infection

110

3.1 Optimisation of experimental conditions for infection and SGs detection 111

3.2 Chemical induction of SGs using oxidative stress in CRFK cells 117

3.3 Optimisation of the markers and conditions used for stress granules detection

in CRFK cells

119

3.4 Optimisation of the markers used for stress granules detection in RAW264.7

cells

123

3.5 Stress granules formation during FCV infection. 125

3.6 Stress granules formation during MNV-1 infection. 134

3.7 FCV infection impairs the assembly of stress granules induced by sodium

arsenite

139

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3.8 FCV infection impairs the assembly of stress granules induced by hydrogen

peroxide

145

3.9 Effect of Stress granules induction on FCV replication 149

3.10 Effect of FCV infection on pre-assembled stress granules 151

3.11 Effect of UV-inactivation on the inhibition of stress granules assembly 156

3.12 Discussion 160

3.12.1 Absence of stress granules in stressed and MNV-infected RAW264.7

macrophages

161

3.12.2 FCV replication impairs stress granules assembly during infection

162

CHAPTER 4 Paracrine induction of Stress Granules by FCV-infected cells 167

4.1 Generation of virus-free cell culture supernatant 168

4.2 Effect of FCV-infected cells supernatant on Stress granule formation 171

4.3 Effect of interferon α on stress granule formation in CRFK cells 174

4.4 Effect of Ribonuclease A (RNase A) on the ability of FCV-infected cells

supernatant to induce stress granule assembly

178

4.5 Effect of Heat-Shock on the ability of FCV-infected cells supernatant to

induce stress granule assembly

181

4.6 Effect of FCV-infected cells supernatant on wide range of cells’ ability to

assemble stress granules

185

4.7 Discussion 188

4.7.1 FCV infection leads to paracrine induction of SG assembly 188

4.7.2 Different types of stress granules can be assembled in response to infection 189

4.7.3 Could this paracrine induction of SGs reflect antiviral mechanism? 190

4.7.4 Investigating the nature of the soluble mediator(s) triggering stress granules 192

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assembly

CHAPTER 5 Regulation of P-Bodies formation during FCV and MNV-1 infection

5.1 Optimisation of the markers and conditions used for P-bodies detection 195

5.2 Effect of MNV-1 infection on P-bodies formation 200

5.3 Effect of FCV infection on the assembly of P-bodies 210

5.4 Effect of chemical induction of oxidative stress on P-bodies assembly 213

5.5 Discussion 217

5.5.1 Modulation of P-bodies assembly during MNV-1 and FCV infection 217

6.1 CHAPTER 6 Conclusion 223

7.1 CHAPTER 7 Reference 229

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LIST OF FIGURES

CHAPTER 1

1.1 Proposed model of induction stress granules (SG) assembly by different

conditions and stimuli

13

1.3 Eukaryotic mRNA decay pathways 27

1.3 Components of P-bodies and stress granules 31

1.4 Proposed model for the role of cytoplasmic RNA granules in mRNA fate 34

1.5 Mechanisms of stress granules disruption by viruses 37

1.6 Mechanisms of P-bodies disruption by viruses 47

1.7 Crystal structures of Norwalk Virus, MNV-1 and FCV by cryo-electron

microscopy

50

1.8 Phylogenetic analysis of the capsid amino acid sequence of Norovirus strains 56

1.9 Organization of representative caliciviruses genome 72

1.10 The structure of Norovirus major capsid protein, viral protein 1 (VP1) 77

1.11 Proposed replication mechanism of the Human Norovirus (HuNoV) 79

1.13 Schematic diagram of cap and VPg-dependent translation initiation

82

CHAPTER 2

2.1 Preparation of the FCV-free supernatant from FCV-infected cells for

immunofluorescence studies

104

CHAPTER 3

3.1 Optimization of FCV MOI for immunofluorescence studies 113

3.2 Optimization of MNV-1 MOI for immunofluorescence studies 115

3.3 Optimization of p76 antibody dilution for immunofluorescence studies 116

3.4 Induction of stress granules by oxidative stress 118

3.5 Detection of stress granules by oxidative stress in FCV-infected cells using

various markers

120

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3.6 3.6 Optimization of FBS concentration for detect of stress granules in CRFK

cells

122

3.7 Detection of stress granules in RAW264.7 cells using various markers 124

3.8 Detection of stress granules during FCV infection 126

3.9 Quantification of the number of cells displaying stress granules during FCV

infection

127

3.10 Detection of stress granules during FCV infection using PABP-1 as a marker 129

3.11 Quantification of the number of cells displaying stress granules during FCV

infection using PABP-1

130

3.12 Detection of SGs in FEA cells during FCV infection 131

3.13 Detection of SGs formation during FCV infection using different markers 133

3.14 Detection of stress granules during MNV-1 infection 135

3.15 Detection of stress granules during MNV-1 infection using various markers 136

3.16 Detection of stress granules during MNV-1 infection in BV-2 cells using

different markers

138

3.17 Detection of stress granules induced by NaAsO2 following FCV infection 140

3.18 Quantification of the number of cells displaying stress granules induced by

NaAsO2 following FCV infection at different time points

141

3.19 Detection of stress granules induced by sodium arsenate (SA) following FCV

infection using PABP-1 as a marker

143

3.20 Quantification of the number of cells displaying stress granules induced by

sodium arsenate following FCV infection using PABP-1 as a marker

144

3.21 Detection of stress granules induced by hydrogen peroxide (H2O2) following

FCV infection

147

3.22 Quantification of the number of cells displaying stress granules induced by 148

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hydrogen peroxide (H2O2)

3.23 Effect of stress granules induction on FCV replication 150

3.24 Effect of FCV infection on pre-assembled stress granules 154

3.25 Quantification of the number of cells displaying stress granules upon sodium

arsenate (SA) treatment and FCV infection

155

3.26 Effect of UV-inactivation on FCV replication 157

3.27 Effect of UV-inactivation on the inhibition of Stress Granules assembly 158

3.23 Quantification of the number of cells displaying stress granules during FCV or

UV-inactivated FCV infection and sodium arsenate (SA) treatment

159

CHAPTER 4

4.1 Detect of FCV replication by TCID50 following virus elimination 170

4.2 Induction of stress granules by cell supernatant 172

4.3 Quantification of the number of cells displaying stress granules following

treatment with cell supernatant

173

4.4 Effect of interferon treatment on stress granule formation 176

4.5 Quantification of the number of cells displaying stress granules following

treatment with IFNα

177

4.6 Effect of RNase A treatment on stress granule formation 179

4.7 Quantification of the number of cells displaying stress granules following

treatment with RNase A

180

4.8 Effect of heat shock on stress granule formation 183

4.9 Quantification of the number of cells displaying stress granules following

treatment with heat shocked supernatant

184

4.10 Effect of FCV-infected cells supernatant on wide range of cells 186

4.11 Quantification of the number of cells displaying stress granules following 187

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treatment wide range of cells with FCV-infected cell supernatant

CHAPTER 5

5.1 Optimization of Dcp1 and Xrn1 antibody dilution in CRFK cells for

immunofluorescence studies

196

5.2 Optimization of Dcp1 and Xrn1 antibody dilution in RAW264.7 cells for

immunofluorescence studies

197

5.3 Optimization of foetal bovine serum concentration for detect of P-bodies in

CRFK cells

199

5.4 Detection of P-bodies during MNV-1 infection 202

5.5 Quantification of the number of cells displaying P-bodies during MNV-1

infection

203

5.6 Detection of P-bodies during MNV-1 infection using BV-2 cells 205

5.7 Quantification of the number of cells displaying P-bodies during MNV-1

infection in BV-2 cells

206

5.8 Detection of P-bodies during MNV-1 infection using Xrn1 as a marker 209

5.9 Detection of P-bodies during FCV infection 211

5.10 Quantification of the number of cells displaying P-bodies during FCV infection 212

5.11 Effect of the sodium arsenite and hydrogen peroxide treatment on P-body

assembly

215

5.12 Quantification of the number of cells displaying P-bodies upon treatment with

the sodium arsenite and hydrogen peroxide

216

CHAPTER 6

6.1 Proposed working model for the effect of FCV and MNV-1 on stress granules

and P-bodies.

228

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LIST OF TABLES

CHAPTER 1

1.1 Components of stress granules 8

1.2 Components of P-bodies 22

1.3 Taxonomy of the Caliciviridae family

49

CHAPTER 2

2.1 Primary and secondary antibodies that are used in immunofluorescence studies 99

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LIST OF ABBREVIATIONS

2Apro 2Apro protease

3Cpro 3C-like Cysteine protease

4E-BP eIF4E-binding protein

40S Eukaryotic small ribosomal subunit (40 Svedberg)

60S Eukaryotic larg ribosomal subunit (60 Svedberg)

APOBEC3G Apolipoprotein B mRNA-editing enzyme, catalytic polypeptide-like 3G

APOBEC3F Apolipoprotein B mRNA-editing enzyme, catalytic polypeptide-like 3F

ADAR Adenosine deaminase, RNA-specific

Ago2 Argonaute RISC catalytic component 2

ALS amyotrophic lateral sclerosis

ATF4 Activating transcription factor 4

ATP Adenosine triphosphate

BNoV Bovine norovirus

BRF1 Butyrate response factor 1

Caprin-1 Cell cycle associated protein 1

CRFK Crandell Rees Feline Kidney cells

CIRP Cold-inducible RNA-binding protein

CPEB Cytoplasmic polyadenylation element binding protein

CrPV Cricket paralysis virus

CVB3 coxsackievirus B3

GADD34 DNA damage–inducible protein 34

DCP1 Decapping mRNA 1

DDX3 DEAD box helicase 3

DDX6/RCK DEAD box helicase 6

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DMEM Dulbecco’s Minimum Essential Medium

DMSO Dimethyl sulfoxide

DV Dengue virus

EBHSV European Brown Hare Syndrome Virus

eEF2 eukaryotic elongation factor 2

Edc1-3 Enhancer of decapping protein 1-3

EDC4/GE-1/Hedls Enhancer of mRNA decapping 4

EMCV encephalomyocarditis virus

eIF Eukaryotic initiation factor

eIF1A Eukaryotic translation initiation factor 1A

eIF2 Eukaryotic translation initiation factor 2

eIF3 Eukaryotic translation initiation factor 3

eIF4A Eukaryotic translation initiation factor 4A

eIF4B Eukaryotic translation initiation factor 4B

eIF4G Eukaryotic translation initiation factor 4G

eIF4H Eukaryotic translation initiation factor 4H

eIF5B Eukaryotic translation initiation factor 5B

EMCV Encephalomyocarditis virus

EV71 Enterovirus 71

FAST Fas-activated Ser/Thr kinase

FBS Fœtal Bovine Serum

FCV Feline calcivirus

FCV-VSD FCV-associated virulent systemic disease

FEA Feline Embryonic Airway

FMDV Foot-and-mouth disease virus

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FMR Fragile X mental retardation

FAK focal adhesion kinase

FRMP Fragile X mental retardation protein

FUS Fused in Sarcoma

FXS Fragile X syndrome

O-GalNAc O-N-acetylgalactosamine

G3BP1 Ras-GTPase-activating protein SH3-domain-binding protein 1

GCN2 General control non-derepressible 2

GDP Guanosine diphosphate

gRNA Genomic RNA

Grb7 growth factor receptor-bound protein 7

GTP Guanosine triphosphate

GW182/ TNRC6A GWbodies 182/ Trinucleotide repeat-containing 6A

(HBGAs) Histo-blood group antigens

HCV Hepatitis C virus

HDAC6 Histone deacetylase 6

HIV-1 human immunodeficiency virus 1

HLTV-1 human T-cell leukaemia virus type 1

HRI Heme-regulated eIF2a kinase

HIV Human Immunodeficiency Virus

hpi Hour post infection

HSGs Heat shock granules

HSP Heat shock protein

HSV-1or 2 herpes simplex virus-1 or 2

HuNoV Human Norovirus

xviii

HuR/ELAVL1 Hu antigen R/ELAV-like RNA-binding protein 1

IAV influenza A virus

ICTV International Committee on Taxonomy of Viruses

IP5K Ins Intracellular localization of human Ins(1,3,4,5,6)P5 2-kinase

IFN Interferon

JAM-A Junctional Adhesion Molecule A

JEV Japanese encephalitis virus

JUNV Junín virus

KSHV Kaposi’s Sarcoma-associated Herpesvirus

kDa Kilo Dalton

LaVs Lagoviruses

MBNL1 Muscleblind-like protein 1

MCPIP1 Monocyte chemotactic protein-induced protein 1

MEF Mouse embryo fibroblasts

MEM Eagle’s Minimum Essential Medium

(MHE) Mouse hepatitis coronavirus

miRNA Micro RNA

MLN51 Metastatic lymph node 51

MNV Murine Norovirus

MOI Multiplicity of Infection

mRNA messenger RNA

mRNPs messenger ribonucleoproteins

MRV mammalian orthoreovirus

mTOR Mammalian target of rapomycin

M7GpppG 7-methyl guanosine cap structure of RNA

xix

NeVs Neboviruses

NHS UK National Health Service

NMD Nonsense-mediated decay

NoVs Noroviruses

NS Non-structural protein

nt nucleotide

NV Norwalk Virus

ORF Open reading frame

PABP Poly (A)-Binding Protein

Pat A Pateamine A

PCBP2 Poly (rC)-binding protein

PKR Protein kinase R

PEC porcine enteric calicivirus

PERK Protein kinase RNA-like endoplasmic reticulum kinase

PSaV Porcine SaV

PV Poliovirus

PIC Pre-initiation complex

Poly (A) Polyadenylation tail

PTB Polypyrimidine tract-binding protein

RACK1 The Receptor for Activated C Kinase 1

RAG2 Mice lacking recombination-activation gene 2

RAP55/LSM14A RNA-associated protein 55

RdRp RNA dependent RNA polymerase

RHDV Rabbit Haemorrhagic Disease Virus

RISC RNA-induced silencing complexes

xx

Rpb4 RNA polymerase II subunit B4

RRL Rabbit Reticulocyte Lysate

RSV Respiratory syncytial virus

RT Room Temperature

rVLPs Recombinant virus like-particles

SaVs Sapoviruses

SDS-PAGE Sodium dodecyl sulphate polyacrylamide gel electrophoresis

SFV Semliki Forest Virus

sgRNA Sub-genomic RNA

siRNA Small Interfering RNA

SMN Survival of motor neuron

SMSLV San Miguel Sea Lion Virus

STAT 1 Signal Transducers and Activators of Transcription

TDP-43 TAR DNA-binding protein 43

TCID50 50% Tissue Culture Infectious Dose

TC Ternary complex

TH2RNP TIAR-HIV-2 ribonucleoprotein

TIA1 T cell restricted intracellular antigen-1

TIAR TIA-1-related protein

TMEV Theiler’s murine encephalomyelitis virus

TRAF2 TNF receptor-associated factor 2

UPF1,2,3 Regulator of nonsense transcripts 1,2,3

USP10 Ubiquitin specific peptidase 10

TTP Tristetraprolin

TV Tulane virus

xxi

UTR Untranslated region

UV Ultra-violet

VeVs Vesiviruses

VLP Virus-Like Particle

VP Virion Protein

VPg Viral Protein Genome-linked

VSV Vesicular Stomatitis Virus

VESV Vesicular Exanthema of Swine Virus

vhs Viral host shutoff protein

wt Wild Type

WHO Word Health Organisation

WNV West Nile virus

Xrn1 5’-3’ Exoribonuclease 1

ZBP1 Z-DNA-binding protein 1

1

Chapter 1

Introduction

2

1.1 RNA Granules

RNA granules are self-assembling structures, or foci, that form in response to stress

conditions via the aggregation of non-translating mRNAs with several proteins to form

messenger ribonucleoproteins (mRNPs) granules (Anderson & Kedersha, 2006; Buchan,

2014; Reineke & Lloyd, 2013; Tsai & Lloyd, 2014). The assembly of these membrane-less

granules occurs either in the nucleus (nuclear mRNPs granules) or in the cytoplasm

(cytoplasmic mRNPs granules) and both are found in all type of eukaryotic cells, contributing

to many events of mRNA metabolism and processing (such as translation, subcellular

localization, turnover) (Balagopal & Parker, 2009; Buchan, 2014; Reineke & Lloyd, 2013).

Nuclear granules including Cajal bodies, nuclear speckles, Polycomb bodies,

paraspeckles, histone locus bodies and nuclear stress bodies are involved in the modulation of

several nuclear activities such as RNA splicing, modification, assembly and storage of small

nuclear RNP (snRNPs), mRNA maturation and export, histone mRNA transcription and

processing, gene transcription and others (Anderson & Kedersha, 2009a; Caudron-Herger &

Rippe, 2012; Mao et al., 2011). The cytoplasmic RNA granules are classified according to

cellular context, marker proteins, and function into various types including stress granules

(SGs), Processing-bodies (P-bodies; PBs) in somatic cells, germ granules in germ cells and

neuronal granules in neurons (Buchan, 2014). These granules have been implicated in innate

immunity, post-transcriptional regulation of gene expression, mRNA storage and mRNA

decay or degradation (Anderson & Kedersha, 2006).

Germ granules of various classes are found in germ cells dependent on their

development stage including nuage (germ cell), sponge, balbiani, and chromatid bodies

(gametes) and germ plasma (embryos). They contain several proteins such as RNA helicase

Vasa involved during germ cell development events such as regulation, localisation, stability,

3

translation and storage of mRNAs (Voronina et al., 2011). In addition, some proteins

involved in mRNA translation such as eukaryotic initiation factor 4E (eIF4E) (Amiri et al.,

2001), and RCK (Navarro & Blackwell, 2005), accumulate in germ granules, as well as

proteins involved in mRNA degradation including Sm/Lsm proteins (Audhya et al., 2005;

Boag et al., 2005), and Decapping proteins 1 and 2 (Dcp1/2) (Lall et al., 2005).

Neuronal RNA granules, also termed transport granules, are mainly involved in

mRNA transport but also play a possible role in mRNA localisation (Anderson & Kedersha,

2006; Krichevsky & Kosik, 2001). In addition to proteins such as Staufen (Thomas et al.,

2005), G3BP1 and HuR/D (Atlas et al., 2004), these granules contain ribosomal subunits

(40S and 60S) and translation initiation factors (eIF2, eIF3 and eIF4E) (Kiebler & Bassell,

2006; Krichevsky & Kosik, 2001).

The most common cytoplasmic granules associated with mRNA turnover are SGs and

PBs. SGs contain non-translating mRNA along with 40S ribosomal subunits as well as a

subset of translation initiation factors and RNA binding proteins, while PBs are enriched with

proteins involved in mRNA degradation and decay (Jain & Parker, 2013a). All these types of

RNA granules contain non-translated mRNA and share many RNA binding proteins which

may reflect the possible relationship between these structures (Buchan, 2014).

1.2 Cytoplasmic mRNPs granules (Stress granules and P-bodies)

In order to maintain cellular homeostasis during changing conditions caused, for

example, by stress, somatic cells apply a variety of regulatory mechanisms allowing the

control of gene expression. One of these mechanisms is the global inhibition or

reprogramming of mRNA translation in response to stress conditions; this impairs gene

4

expression and contributes to mRNA turnover until stress recovery (Shenton et al., 2006).

Cytoplasmic SGs and PBs form in response to translational arrest via the aggregation of non-

translated mRNAs with a variety of proteins, they play critical roles in regulating gene

expression via the control of mRNA translation and degradation during stressful conditions

such as viral infections (Anderson & Kedersha, 2002; Brengues et al., 2005; Decker &

Parker, 2012; Parker & Sheth, 2007). In eukaryotic cells, cytosolic mRNAs are in dynamic

equilibrium among polysomes, where mRNAs are translated, SGs, in which untranslated

mRNAs and associated proteins are stored during conditions of stress, and PBs, which

contain the decay machinery for degrading non-translated mRNA (Anderson & Kedersha,

2006; Parker & Sheth, 2007). The movement of mRNA between these different functional

states may support the roles of these subcellular structures in regulating gene expression

(Brengues et al., 2005; Buchan et al., 2008; Decker & Parker, 2012; Kedersha et al., 2005;

Mollet et al., 2008). Despite the differences between PBs and SGs in composition and

function, both are induced by various types of environmental stress such as heat shock, UV

light, starvation, oxidative stress (arsenite poisoning), and endoplasmic reticulum stress, as

well as under certain physiological conditions and during viral infections (Kedersha et al.,

1999; Towers et al., 2011; Tsai & Lloyd, 2014). Furthermore, they share many components

as detailed in the next section (Buchan, 2014; Kedersha & Anderson, 2009).

1.3 Stress granules (SGs)

SGs were discovered in 1983 as foci, or spherical granules, containing a specific

subset of cellular mRNAs within the cytoplasm of Peruvian tomato (Lysopersicon

peruvianum) cells subjected to heat shock in vitro (Nover et al., 1983). The formation of

these foci was associated with the inhibition of translation in stressed tomato cells and these

5

granules termed heat shock granules (HSGs), containing a 27-kDa heat shock protein

(HSP27) as well as non-translated mRNAs only translated after recovery from stress (Nover

et al., 1983; 1989). This suggested that the formation of HSGs protected the mRNA from the

damage caused by stress conditions, storing it until recovery to enter translation again. Later,

these similar granules containing heat shock protein HSP28 were also observed in the

cytoplasm of heat-shocked mammalian cells (Arrigo, 1990; Arrigo et al., 1988).

Subsequently, several studies have demonstrated that SGs are formed in many types of plant

and animal cells in response to a variety of stress factors including radiation (Moeller et al.,

2004), hypoxia (Gardner, 2008; van der Laan et al., 2012), oxidative stress (Emara et al.,

2012; Kedersha et al., 1999), osmotic stress (Dewey et al., 2011), nutrient stress (Tattoli et

al., 2012), some physiological conditions (Towers et al., 2011), chemical inhibitors of

translation initiation factors (Fournier et al., 2010; Mazroui et al., 2006) and during viral

infections (Lloyd, 2012). Kedersha et al. (1999) demonstrated using mammalian cells that the

RNA-binding proteins T-cell intracellular antigen 1 (TIA-1) and T-cell restricted intracellular

antigen-related protein (TIAR) colocalize with poly (A)-binding protein (PABP) in SGs

formed in response to environmental stress and suggested that the assembly of SGs depends

on the phosphorylation of eukaryotic initiation factor 2α (eIF2 which is enhanced by TIA-1

(Kedersha et al., 1999).

In mammalian cells, studies using cochlear hair cells identified that aminoglycoside

treatments using gentamycin induce TIAR translocation from the nucleus and accumulation

as punctate granules in the cytoplasm (Mangiardi et al., 2004). Moreover, the formation of

Caprin 1-containing stress granules in cochlear hair cells following aminoglycoside-induced

damage was also reported (Towers et al., 2011). Furthermore, the assembly of SGs was found

to be associated with Fragile X mental retardation (FMR) or Fragile X syndrome (FXS), and

the FMR protein (FMRP) which associated with polysomes in normal cells but in stressed

6

cells was found to colocalize with non-translated mRNA in SGs (Kim et al., 2006; Mazroui

et al., 2002). These studies have revealed that the inhibition of protein synthesis is critical for

the induction of SGs formation (Kedersha & Anderson, 2007). Chemical agents that induce

translation termination such as puromycin promote SGs formation (Kedersha et al., 2000).

Supporting this, chemical agents that arrest translation elongation and prevent polysome

disassembly, such as emetine and cycloheximide, also block SGs assembly (Kedersha et al.,

2000).

An increasing number of RNA-binding proteins have been identified as components

of stress granules using immunofluorescence studies, but the exact composition of stress

granules remains unknown and varies according to the stress, cell type or experimental

system (Anderson & Kedersha, 2008; 2009b; Buchan & Parker, 2009; Thomas et al., 2011).

For example, in yeast cells, the SGs induced under acute stress are similar in composition to

mammalian SGs. In contrast, yeast cells exposed to glucose starvation form SGs that are

distinct from mammalian SGs, lacking 40S ribosomal subunits and eIF3 (Buchan et al., 2008;

Hoyle et al., 2007). In Saccharomyces cerevisiae, NaN3-induced SGs contain the initiation

factors eIF3, eIF4A/B, eIF5B and eIF1A, whereas SGs induced by glucose deprivation do not

(Buchan et al., 2011).

1.3.1 Stress granule composition

SGs are formed within 15-20 min of stress exposure by the rapid accumulation of

transient mRNP aggregates, in the cytoplasm of cells responding to stress conditions. In

addition to non-translated poly-adenylated mRNAs, in a complex with PABP, SGs contain

translation initiation factors and a large numbers of RNA-binding proteins involved in

various functions ranging from translation, antiviral response to RNA localisation (Kedersha

7

& Anderson, 2009) (Table 1). Typically, SGs contain non-translated mRNA, 40S ribosomal

subunits (but not 60S), the eIF4F complex (eIF4E, eIF4G, eIF4A) and eIF4B, Poly(A)

binding protein (PABP), eIF3, and eIF2 (Anderson & Kedersha, 2006; Kedersha et al., 2000;

Kedersha et al., 1999; Kimball et al., 2003; Low et al., 2005; Mazroui et al., 2006). The most

characterized RNA-binding proteins are G3BP1 and TIA-I/TIAR which possessing self-

interacting domains that promote SGs assembly (Gilks et al., 2004; Tourriere et al., 2003)

and many other RNA-binding proteins regulating mRNA structure and functions. These

include proteins involved in mRNA silencing such as Argonaute proteins (Ago2 and Ago2)

(Leung et al., 2006), pumilio 2, Rap55, Staufen, TIA-1 and TIA-1-related protein (TIAR)

(Anderson & Kedersha, 2008). Proteins contributing to the antiviral response, including

APOBEC3G and APOBEC3F, are also detected in SGs supporting the role of SGs in defence

against viral infection (Gallois-Montbrun et al., 2007; Kozak et al., 2006). In addition to

translation factors, other proteins associated with mRNA translation are reported as SG

components such as Ataxin-2 (Nonhoff et al., 2007), FAST (Kedersha et al., 2005), FMRP

and FXR1 (Antar et al., 2005). Proteins involved in metabolic signalling pathways such as

G3BP1 (Tourriere et al., 2003), IP5K (Brehm et al., 2007) and TRAF2 (Kim et al., 2005),

have also been detected. Proteins involved in mRNA decay (p54/RCK/DDX6, TTP, BRF-1,

Xrn1, and Dcp1/2) (Kedersha et al., 2005; Rothe et al., 2006; Stoecklin et al., 2002;

Wilczynska et al., 2005), protein enhancing mRNA stability (HuR) (Gallouzi et al., 2000),

mRNA localisation proteins (ZBP1) (Stohr et al., 2006), and proteins important for

transcription (Rpb4 and SRC3) (Lotan et al., 2005), or splicing (MLN51) (Baguet et al.,

2007), have also been found to act as SG components. This evidence regarding the wide

nature of SG components may reflect the many possible roles for these granules in mRNA

metabolism or life cycle (Anderson et al., 2014; Buchan, 2014).

8

Table 1.1 Components of stress granules

Protein Known name Function

Argonaut proteins Ago1 Gene silencing

Ago2 siRNA , slicer

Apolipoprotein B mRNA editing enzyme,

catalytic polypeptide-like 3G

APOBEC3G Cytidine deaminase , antiviral

Cytoplasmic polyadenylation element-

binding protein 1

CPEB1 Translational silencer

Eukaryotic initiation factor 4E eIF4E Translation initiation

Fas-activated serine/threonine

phosphoprotein

FAST Signalling

Human antigen R HuR Stability, splicing

Importin 8 Imp8 Shuttling, silencing

Lin28 Lin28 Block let-7 processing

LINE 1 ORF1p

Sm-like protein1 Lsm1 mRNA splicing

hmex-3B hmex-3B Germline development

Musashi Musashi translational control that regulate

multiple stem cell populations

DEAD box protein RCK/p54 RCK/ p54 Translation /decay

RNA-associated protein 55 RAP55/Lsm14 shuttling

Roquin proteins Roquin E3 ligase, promote mRNA

degradation by recruiting the

Ccr4-Caf1-Not deadenylase

complex

Smaug protein Smg Translational silencer

T-cell intracellular antigen 1/ T-cell

restricted intracellular antigen-related

protein

TIA-1/TIAR Splicing, translational silencer

Tristetraprolin/Butyrate response factors

1

TTP/BRF-1 mRNA decay

Exoribonuclease Xrn1 mRNA decay (5' 3' exonuclease

Y box binding protein 1 YB-1 mRNA chaperone

A-kinase anchor protein 350A AKAP350A Protein scaffold

apolipoprotein B mRNA-editing enzyme

1

APOBEC1 RNA editing

Calreticulin calregulin,

CRP55, CaBP3,

Cell division cycle and apoptosis

regulator protein 1

CCAR1 Transcriptional co-activator

Caprin-1 Caprin-1 Cell cycle

Cell division cycle and apoptosis

regulator protein 1

CCAR1 Transcriptional co-activator

Caprin-1 Caprin-1 Cell cycle

Cold-inducible RNA-binding protein CIRBP Translational silencer

Death-associated protein 5 DAP5/p97/NAT1 Translation

9

Protein Known name Function

DEAD box protein A DBPA Splicing

ATP-dependent RNA helicase 1 DDX1 ds DNA breaks

ATP-dependent RNA helicase 3 DDX3 RNA helicase

Eukaryotic initiation factor 1 eIF1 Translation initiation

Eukaryotic initiation factor 2 α eIF2α Translation initiation

Eukaryotic initiation factor 3 eIF3 Translation initiation

Eukaryotic initiation factor 4A,B eIF4A, eIF4B Translation initiation

Eukaryotic initiation factor 4G eIF4G Translation initiation

Eukaryotic initiation factor 4H eIF4H Translation initiation

Ewing sarcoma protein EWS oncogene

Focal Adhesion Kinase FAK Motility, signalling

Fragile mental retardation protein / Fragile

X mental retardation syndrome-related

protein 1

FMRP/FXR1 Translation, splicing,

microRNA

Fused in Sarcoma FUS Motility

Folate-binding protein /KH-type splicing

regulatory protein

FBP/KSRP mRNA decay, miRNA

processing

Growth factor receptor-bound protein 7 GRB7 Protein scaffold

Ras GTPase-activating protein-binding

protein

G3BP Helicase, protein scaffold

Histone deacetylase 6 HDAC6 Deacetylase stress signalling

heterogeneous nuclear ribonucleoprotein A1 hnRNPA1 Splicing , multifunctional

heterogeneous nuclear ribonucleoprotein K hnRNPK mRNA processing

heterogeneous nuclear ribonucleoprotein Q hnRNPQ Splicing

Heat shock protein 27 HSP27 Heat shock

Heat shock protein 90 HSP90 Molecular chaperon

Intracellular localization of human

Ins(1,3,4,5,6)P5 2-kinase

IP5K Ins Protein scaffold

Muscleblind-like protein 1 MBNL1 Alternative splicing

Metastatic lymph node 51 MLN51 Splicing

Ploy A binding protein PABP Translation

Protein activator of the interferon-induced

protein kinase

PACT RISC, PKR activator

Plasminogen activator inhibitor 1 RNA-

binding protein

PAIRBP1

Plakophillin1/3 PKP1/3 Adhesion

plasma membrane-related Ca(2+)-ATPase 1 PMR1 Endonuclease

Pumilio 2 Pumilio 2 Development

The Receptor for Activated C Kinase 1 RACK1 Signalling, polarity

RNA binding motif protein 42 RBM42

RNA Helicase associated with AU-rich

element

RHAU helicase RNA helicase

Ribosomal s6 kinase2 RSK2 Kinase

Ribonuclease P protein subunit p20 Rpp20 RNAse P supunit

RNA polymerase II subunit B4 Rpb4 mRNA transcription

Src-associated in mitosis 68 kD SAM68 Multifunction

Stress Granule/Nucleolar Protein

SCNP Ribosome maturation

10

Protein Known name Function

The nuclear receptor coactivator 3 SRC3 Transcriptional co-activator

Staufen Staufen ds RNA binding

Survival of Motor Neuron SMN snRNP assembly

TATA-binding protein-associated factor 2N TAF15 Oncogene

Tudor domain-containing protein 3 TUDOR-3

Translin TSN Antiviral nuclease

Ubiquitin Ubiquitin Signalling protein

Z-DNA-binding protein 1 ZBP1 (zip code BP) mRNA localisation,

antiviral

The Tudor-SN protein TSN Transcription,

The energy sensor AMP-activated protein

kinase (AMPK)-2α

AMPK-α2 a central regulator of energy

homeostasis in eukaryotes,

autophagy, cell growth and

polarity to cytoskeletal

dynamics

Ataxin-2 Ataxin-2 Translation

11

1.3.2 Stress granule assembly/disassembly

Responding to stress-induced damage, eukaryotic cells activate mechanisms to save

energy and repair this damage by repressing the translation of house-keeping gene transcripts

which accumulate in SGs, and reciprocally up-regulating the translation of proteins required

to repair the cellular insults (Anderson & Kedersha, 2008). To date, the mechanism by which

stress granules assemble/disassemble is not yet fully understood, and more than 100 genes are

involved in this complex process with over 80 proteins that localise to SGs (Buchan &

Parker, 2009; Ohn et al., 2008). Many factors regulate and promote SG assembly such as

non-translating mRNAs, post-translational modifications, protein-protein interactions, and the

microtubule network (Buchan & Parker, 2009).

The first stage of SG formation is always associated with a shut-off of protein

synthesis that can be caused by different pathways (Kedersha et al., 1999; Mazroui et al.,

2006). Studies have demonstrated that one of the major mechanisms for triggering SG

formation is the phosphorylation of the alpha subunit of eIF2 on serine residue 51 (Holcik &

Sonenberg, 2005; Kedersha et al., 1999). This phosphorylation is carried out by one of four

serine/threonine kinases activated in response to different types of stress (Holcik &

Sonenberg, 2005)(Figure 1.1). The double-stranded RNA-dependent protein kinase (PKR) is

commonly activated by RNA viruses as a part of the interferon antiviral response of cell

(Maggi et al., 2000), but PKR also senses heat shock, oxidative stress and UV irradiation

(Williams et al., 2001). General control non-de-repressible-2 kinase (GCN2) senses amino

acid deprivation and the decrease of other types of nutrients, as well as UV irradiation

(Berlanga et al., 1999). PKR-like endoplasmic reticulum kinase (PERK) responds to the ER

stress caused by the accumulation of unfolded or misfolded proteins in the ER (Harding et

al., 2000; Shi et al., 1998), or hypoxia sensing (Harding et al., 2000). The heme-regulated

inhibitor kinase (HRI) is activated under conditions of intracellular iron deficiency but also

12

senses oxidative stress and heat shock (Han et al., 2001; Lu et al., 2001; McEwen et al.,

2005). Activation of all of these kinases leads to phosphorylation of eIF2, thereby

increasing the affinity (150-fold) of eIF2 for eIF2B, the eIF2 guanine nucleotide exchange

factor, blocking the guanosine diphosphate/guanosine triphosphate GDP/GTP exchange and

formation of the eIF2-GTP-tRNAMet ternary complex (TC). This prevents assembly of the

48S pre-initiation complex and results in the repression of translation initiation (Holcik &

Sonenberg, 2005; Srivastava et al., 1998).

13

Figure 1.1 Proposed model of inducting stress granules (SG) assembly by different conditions

and stimuli. The most characterised induction pathway leading to SGs assembly is the

phosphorylation of eukaryotic translation initiation factor 2α (eIF2α) by the kinases PKR, PERK, and

HRI and GCN2. This blocks the formation of eIF2-GTP-Met-tRNAiMet

ternary complex (TC) and in

turn inhibits translation initiation to trigger the assembly of SGs. In addition SGs formation can be

induced by the alteration of expression level or function of translation factors such as eIF4A, eIF4B,

eIF4H, and poly (A)-binding protein (PABP).

14

The formation of SGs can also be induced independently of the eIF2

phosphorylation pathway via cleavage or alteration, or modification of expression level and

activity, of proteins involved in translation initiation such as eIF4A, eIF4G (Anderson &

Kedersha, 2009a; Bordeleau et al., 2006; Mazroui et al., 2006), eIF4B, eIF4H, as well as

PABP (Mazroui et al., 2006). The activation or overexpression of key SG–nucleating

proteins such as G3BP1, TIA1, TIAR, TDP-43, Caprin-1, CIRP and CBEP1 also triggers the

assembly of SGs without any additional stimulus (De Leeuw et al., 2007; Gilks et al., 2004;

Kedersha et al., 2013; Tourriere et al., 2003; Wilczynska et al., 2005). However, in most

cases, it is the phosphorylation of eIF2 that leads to a rapid dissociation of polysomes even

if other routes exist (Mazroui et al., 2006).

Therefore, the first step of SG formation can occur via both eIF2 phosphorylation-

dependent and independent pathways that affect translation initiation and result in polysome

disassembly whereupon the cell begins to develop foci to store the stalled translation

initiation complexes (48S mRNPs) (Kedersha et al., 1999). Locally, RNA-binding proteins

also localise to these small foci by their affinity for the suddenly high concentration of free

mRNA in the cytoplasm (primary aggregation) (Bounedjah et al., 2014; Kedersha &

Anderson, 2009). Several factors will mediate the subsequent stages of SG assembly

mediated by proteins containing self-interaction domains that mediate protein-protein or

protein-RNA interactions (Gilks et al., 2004; Kedersha & Anderson, 2002; Tourriere et al.,

2003). Such domains are found in several RNA-binding proteins. For example, TIA-1 and

TIAR contain glutamine and asparagine-rich prion-like domains mediating SG formation via

their self-aggregating ability (Gilks et al., 2004; Kedersha & Anderson, 2002). G3BP1 also

harbours the self-interaction (polyglutamine and asparagine rich) domains, playing a critical

role in SGs assembly (Tourriere et al., 2003), and knockdown of these proteins leads to a

reduced assembly of SGs (Ghisolfi et al., 2012). PABP, which associates with most

15

transcripts, also mediates the aggregation of SGs via its C-terminal domain (Kedersha &

Anderson, 2009). Other RBPs that promote SGs assembly include FMRP/FXRI (Mazroui et

al., 2002), Staufen (Martel et al., 2010), SMN (Hua & Zhou, 2004), TTP (Stoecklin et al.,

2004), BRF1 (Kedersha et al., 2005), and CPEB (Wilczynska et al., 2005). Proteins lacking

RNA-binding features can also contribute to this process via protein-protein interaction with

certain mRNA binding proteins in SGs, so-called piggyback proteins (Kedersha & Anderson,

2009). For example, SRC3, FAST, PMRI are localized to SGs via interaction with TIA-1

(Yang et al., 2006b; Yu et al., 2007). In addition, TRAF2 and plakophillin 3 that bind to

eIF4G and G3BP1, respectively, are recruited to SGs in a piggyback manner (Hofmann et al.,

2006; Kim et al., 2005). The recruitment of specific SGs proteins components during this

stage is largely associated with an excess of free mRNA (non-polysomal mRNA) and the

nucleic acid/proteins imbalance may also regulate the SGs assembly (Bounedjah et al., 2014).

Protein modifications are also involved in the assembly of SGs by enhancing the

recruitment of several proteins to SGs (Ohn & Anderson, 2010). Phosphorylation and

dephosphorylation of several proteins play a critical role in SGs assembly, for example the

phosphorylation of eIF2 plays a key role in initiation of SG formation (Kedersha et al.,

1999). The overexpression and phosphorylation of G3BP also regulate SG

assembly/disassembly, while the overexpression of G3BP1 induces SGs assembly; the

phosphorylation of this protein at serine residue 149 inhibits SGs formation (Tourriere et al.,

2003). Phosphorylation of other proteins has also been reported to regulate SGs

assembly/disassembly such as phosphorylation of growth factor receptor-bound protein 7

(Grb7) by focal adhesion kinase (FAK) in heat shock–stressed cells (Tsai et al., 2008).

Phosphorylation of tristetraprolin (TTP) regulates the interaction between SGs and PBs

(Stoecklin et al., 2004), and phosphorylation of eIF4E-transporter (4E-T) enhances stress-

dependent PBs assembly (Cargnello et al., 2012). Protein modification such as the

16

hypusination of eIF5A via a polyamine biosynthetic pathway inhibits SGs assembly (Li et al.,

2010). Moreover, studies have reported that the methylation of the RGC domain of FMRP

and cold inducible RNA-binding protein (CIRP) are required for SGs assembly (De Leeuw et

al., 2007; Dolzhanskaya et al., 2006). Furthermore, Ohn et al., (2008) observed that the O-

GlcNAc modification of ribosomal proteins correlates with SGs formation (Ohn et al., 2008).

Finally, the cytoskeleton and associated motor proteins also participate in

assembly/disassembly of SGs by transporting stalled mRNPs to sites of RNA granule

assembly (Bartoli et al., 2011; Loschi et al., 2009). The depolymerisation of microtubules

using nocodazole or vinblastine prevented the assembly of SGs in a mammalian CV-1 cell

line that was treated with arsenite, while microtubule-stabilizing drugs such as paclitaxel

promote SG formation (Ivanov et al., 2003). It was further proposed that the disruption of

microtubules affects the number and size of SGs rather than inhibiting their formation

(Fujimura et al., 2008; Kolobova et al., 2009; Loschi et al., 2009). In contrast, Kwon et al.,

(2007) reported that the stabilization of microtubules was not associated with SGs assembly

(Kwon et al., 2007). In addition to microtubules, microtubule motor proteins such as dynein

and kinesin localize to SGs and also contribute in SGs assembly/disassembly. Studies have

reported that dynein facilitates SG assembly and the absence of this protein from cells treated

with arsenite has a negative impact on the size and number of SGs (Kwon et al., 2007; Loschi

et al., 2009), while kinesin plays a role in disassembly of SGs after stress recovery (Loschi et

al., 2009). According to these findings, the role of microtubules and associated motor

proteins is to facilitate coalescence of small aggregates to form large aggregates (Kolobova et

al., 2009; Tsai et al., 2009). Furthermore, pre-existing PBs may be required for the assembly

of SGs (Buchan et al., 2008). Depending on the SGs component, the mRNAs that localise to

SGs can either exit SGs and re-enter translation, detach from the eIF3/40S complex and

17

localise to PBs for degradation via their binding to specific proteins such as TTP, or be stored

in SGs for a prolonged time (Kedersha & Anderson, 2009).

The SGs disassembly process begins when the cells recover from the stress naturally

or artificially (using drugs stabilising polysomes) and translation returns to normal (Anderson

& Kedersha, 2008; Nadezhdina et al., 2010). Other factors such as turnover of their protein

component, chaperone proteins, and autophagy are possibly involved in SG disassembly

(Buchan, 2014).

1.3.3 Functions and importance of stress granules

In addition to their role in maintaining cellular homeostasis during noxious conditions

via retention and protection of the specific mRNAs stored in a repressed state enabling the

cell to repair the cellular damage until stress recovery (Hofmann et al., 2012; Kedersha et al.,

2005; Tsai & Lloyd, 2014), SGs might participate in a variety of functions mediated by SG-

associated proteins, which include translation, decay, splicing, localization, stability,

signalling, and response to viral infections (Buchan, 2014; Kedersha & Anderson, 2009).

SG proteins TTP and BRF1/2 that bind to 3′ untranslated regions of mRNA promote

the interaction between SGs and PBs to facilitate mRNA decay within PBs (Anderson &

Kedersha, 2008; Brengues et al., 2005; David Gerecht et al., 2010; Kedersha et al., 2005;

Yang et al., 2006a). Zipcode-binding protein 1 (ZBP1) controls mRNA turnover, translation

and localisation in non-stressed cells (Yisraeli, 2005). During cellular stress, the interaction

of this protein with specific mRNAs within SGs is required for the stabilization of associated

mRNAs, regulating the mRNA fate during stress conditions (Stohr et al., 2006). Moreover,

knockdown of ZBP1 leads to destabilisation of specific mRNAs (Stohr et al., 2006). Several

18

SG proteins, including DBPA, FASTK, FMRP/FXR1, hnRNPA1, HuR, SAM68, TIA-1 and

TIAR shuttle between the nucleus and cytoplasm, and this shuttling might play a role in

alternative splicing and microRNA processing (Kedersha & Anderson, 2009). The temporary

sequestration of signalling proteins such as G3BP1, FXR1/FMRP, TIA-1/TIAR, TRAF2,

RACK1 and FAST into SGs may affect multiple signalling pathways as well as RNA

metabolism, thus regulating the survival of the stressed cell (Anderson & Kedersha, 2008;

Kedersha et al., 2013; Kim et al., 2005). Stress granules might also function as a regulator of

cell death (Mokas et al., 2009). The sequestration of pre-apoptotic proteins such as RCCK1,

ROCK1, and TRAF2, as well as mRNA encoding pro-apoptotic proteins inhibits the

apoptotic response (Arimoto et al., 2008; Kim et al., 2005; Moeller et al., 2004). In addition,

the disruption of RNA granule function is associated with several diseases including

neurodegenerative diseases and dementias, immunological and viral diseases (Bloch et al.,

2013; Buchan, 2014; Lloyd, 2013). These diseases usually correlate with mutation of proteins

found as components of SGs such as FMRP, fused in sarcoma (FUS), transactive response

(TAR) DNA binding protein-43 (TDP-43) and ataxin-2 (Anderson & Kedersha, 2008;

Shelkovnikova et al., 2013). For example, Fragile X syndrome (FXS) results from mutations

within the X-linked FMR1 gene which lead to neuronal cell death and may be correlated with

the inhibition of SG assembly (Didiot et al., 2009; Kim et al., 2006). It was proposed that

impairing SG assembly prevents the reprogramming of SG FMRP protein translation (that

protect the cells from sudden environmental stress) leading to FXS (Didiot et al., 2009; Kim

et al., 2006). Two SG proteins, TDP-43 and FUS/TLS, which harbour prion-like domains,

play critical roles in amyotrophic lateral sclerosis (ALS), a neurodegenerative disease

primarily affecting motor neurons (Cleveland & Rothstein, 2001). Mutations in TDP-43

protein increase the tendency of this protein to form SGs in response to stress as well as its

binding to other proteins within SGs, such as TIA-1 or FUS (Liu-Yesucevitz et al., 2010).

19

The sequestration of these proteins in SGs reduces their level in the nucleus and

neurodegeneration could arise from either low levels of nuclear FUS and TDP-43 or

increased formation of SGs (Anderson et al., 2014). SGs have also been proposed to be

involved in the pathogenesis of other diseases such as spinal motor atrophy (Hua & Zhou,

2004), ischemia–reperfusion (Kayali et al., 2005), Huntington’s chorea and Alzheimer’s

disease (Vanderweyde et al., 2013). SGs and PBs are also involved in the mechanisms used

by cancer cells to re-program gene expression and enhance cell survival by modulating

cellular signalling pathways, metabolic machinery, and stress response programs (Anderson

et al., 2014). Several studies have reported the involvement of RNA granules in cancer

development as well as demonstrating an interplay with the response to chemotherapy and

specific drugs (Anderson et al., 2014). For instance, an association between

chemotherapeutic treatments using bortezomib and increased SGs formation was found to be

necessary for cancer cell survival. Using different types of cancer cell line, including colon

cancer (Caco), cervical cancer (HeLa) and lung cancer (Calu-I) cell lines, formation of SGs

was found to be associated with the ability to resist bortezomib-mediated apoptosis, (Fournier

et al., 2007). The antimetabolite 5-fluorouracil (5-FU) used for treatment of different types of

cancer such as breast, colorectal, head and neck cancer (Longley et al., 2003), also induces

eIF2α phosphorylation-dependent-SGs through activation of PKR leading to cell survival via

the sequestration of RACK1, a protein mediating cell survival and apoptosis (Kaehler et al.,

2014; Longley et al., 2003). Other types of drugs such as Pateamine A (Pat A) (Bordeleau et

al., 2005), hippuristanol (Bordeleau et al., 2006), and silvestrol (Bordeleau et al., 2008),

affecting eIF4A activity and leading to translation initiation inhibition, induce SG assembly

and are anti-tumour agents in many types of cancer (Anderson et al., 2014).

In recent years, there has been growing body of evidence to indicate that SGs and PBs

form part of the cellular response to stress triggered by viral infection (Beckham & Parker,

20

2008; Lloyd, 2013) (see section 1.6). As discussed above, the importance of RNA granules is

not just as a means of storage or degradation of stalled mRNAs, but also because of their

influence on several aspects of cell metabolism and gene regulation during abnormal or

unsuitable conditions (Anderson et al., 2014).

1.4 Processing bodies (P-bodies or PBs)

PBs are the second major type of cytoplasmic RNA granules associated with stressed

or unstressed somatic cells of all eukaryotes such as vertebrates, invertebrates, and plants as

well as in yeast and trypanosomes; and their function is conserved from yeast to human (Jain

& Parker, 2013b; Kedersha & Anderson, 2009; Kulkarni et al., 2010; Parker & Sheth, 2007).

The presence of PBs is constitutive in the cytoplasm of unstressed cells, but their number and

size can increase several fold in response to stress (Anderson et al., 2014; Jain & Parker,

2013b; Kulkarni et al., 2010; Parker & Sheth, 2007; Teixeira et al., 2005). The presence

within PBs of enzymes required for mRNA degradation and surveillance, translational

repression, and RNA silencing might reflect their association with translation repression

and/or mRNA decapping and degradation (Baguet et al., 2007; Brengues et al., 2005; Decker

& Parker, 2012; Decker et al., 2007; Parker & Sheth, 2007). PBs were first observed by

Bashkirov in 1997 as a punctate granular signal of the mammalian Xrn1 exoribonuclease, the

main cytoplasmic 5′→3′ exoribonuclease in eukaryotic cells, in the cytoplasm of a mouse

fibroblast cell line and thus named “Xrn1 foci” (Bashkirov et al., 1997). Then, Eystathioy in

2002 observed a novel protein, GW182, localised to these foci in the cytoplasm of patients

suffering from motor and sensory neuropathy, naming this structures as GW182 bodies

(Eystathioy et al., 2003). Later, other proteins involved in decay mechanism were found to

localise to PBs, including the decapping enzymes Dcp1/Dcp2 (Ingelfinger et al., 2002), the

21

decapping activators Hedls (Yu et al., 2005), Edc3 (Fenger-Gron et al., 2005), Pat1 (Scheller

et al., 2007), Lsm1-7 (Ingelfinger et al., 2002), and RCK (p54, DDX6) (Wilczynska et al.,

2005). Due to the presence of RNA decay machinery components, these cytoplasmic foci are

also called ‘decapping-bodies’ or ‘mRNA decay bodies’, but the most widely accepted name

is P-bodies or PBs (Sheth & Parker, 2003). Moreover, PBs are enriched with other types of

proteins such as RNA degradation proteins, the deadenylase complex CCR4/CAF1/NOT

(Zheng et al., 2008), its enhancer TOB2 (Ezzeddine et al., 2007), nonsense-mediated mRNA

decay proteins UPF1, UPF2, UPF3, SMG5, SMG6 and SMG7, ARE-mediated decay factors

TTP, BRF1 and BRF2 (Franks & Lykke-Andersen, 2007; Kedersha et al., 2005; Stoecklin &

Anderson, 2007), and the RNAi machinery component Argonaut (Sen et al., 2005). In

addition, PBs contain protein involved in translation regulation such as eIF4E (Andrei et al.,

2005), eIF4E-T (Ferraiuolo et al., 2005), FASTK (Kedersha et al., 2005), CPEB (Wilczynska

et al., 2005), and PCBP2 (Fujimura et al., 2008). Table 1.2 shows several other proteins that

can be detected as components of PBs. Moreover, PBs contain translationally repressed

mRNA in response to a variety of stresses such as oxidative stress, UV stress, osmotic shock

and nutritional deprivation (Buchan et al., 2008; Buchan et al., 2011; Izawa et al., 2007;

Teixeira et al., 2005; Yoon et al., 2010).

22

Table 1.2 Components of P-bodies

Protein Known name Function

C-C chemokine receptor type 4 CCR4 Transcription, ribosome

biogenesis

cytoplasmic polyadenylation element binding

protein

CPEB Translation regulator

Decapping enzymes Dcp1and Dcp2 mRNA decay

Enhancer of decapping protein 1,2 Edc1,2 mRNA decay

Enhancer of decapping protein 3 Edc3 mRNA decay

Eukaryotic initiation factor 4E eIF4E Translation initiation

Eukaryotic initiation factor transporter 4E-T eIF4E-T mRNA decay

GW182/Trinucleotide repeat-containing gene

6A protein

GW182/TNRC6B siRNA

Guanylate binding protein 2, interferon-

inducible

Gbp2 mRNA export

Eukaryotic initiation factor 4G eIF4G Translation initiation

Hedls/Ge-1 Hedls/Ge-1 Enhancing mRNA

decapping

Heterogeneous nuclear ribonucleoprotein A3 hnRNPA3 Splicing, multifunctional

Importin-8 Importin-8

Putative helicase MOV-10 MOV10/Armitage RNA-directed transcription

NFX7 NFX7 mRNA export

Protein interacting with APP tail-1

Pat1/Pat1L Decapping

CCR4–CAF1–NOT complex CCR4–CAF1–NOT

complex

Deadenylation

Regulator of nonsense transcripts 1,2,3 UPF1,2,3 Nonsense-mediated decay

(NMD)

Eukaryotic release factor 1 and 3 eRF1 and eRF3 Translation termination

exonuclease Xrn1 5′ to 3′ exonuclease

Rap55 Rap55/Scd6 Translation repressor

Fas activated serine/ threonine

phosphoprotein FAST

Poly C- binding protein 2 PCBP2 Translation regulator

23

1.4.1 P-bodies assembly/disassembly

Assembly of PBs is initiated when translation is repressed in response to a cellular cue

by the interaction of non-translated mRNA with various proteins forming mRNPs, followed

by the aggregation of these mRNPs via self-interaction domains leading to the formation of

PBs visible under microscopy (Decker et al., 2007; Eulalio et al., 2007a). Generally, this

aggregation is reversible, and mRNPs can re-enter translation, or in some cases mRNPs

undergo further rearrangement via association with specific proteins that facilitate the decay

or degradation of mRNA (Bhattacharyya et al., 2006; Brengues et al., 2005; Eulalio et al.,

2007a). Whether the formation of PBs is a de novo process or the mRNA and associated

proteins are localised to pre-existing structures remains an unresolved issue (Jakymiw et al.,

2007).

The mechanism of PB assembly is not entirely understood and varies depending on

the cell system (Decker et al., 2007; Franks & Lykke-Andersen, 2007; 2008). More than 40

genes have been proposed to be involved in the assembly process (Ohn et al., 2008). The

presence of mRNA is required for formation and integrity of PBs (Andrei et al., 2005;

Eystathioy et al., 2002; Teixeira et al., 2005). Treatments with cycloheximide, which traps

the mRNAs in polysomes, or with RNase both lead to the disappearance of visible PBs in

mammalian cells (Cougot et al., 2004; Eulalio et al., 2007a; Franks & Lykke-Andersen,

2008; Teixeira & Parker, 2007). In addition to repressed mRNA, specific PB proteins are

critical for the formation of a small core during PB assembly, and subsequently many other

proteins contribute to this process (Decker et al., 2007; Eulalio et al., 2007a; Kulkarni et al.,

2010; Liu et al., 2005; Yu et al., 2005).

24

In yeast, three proteins are essential for assembling PBs: Edc3 (Decker et al., 2007;

Kulkarni et al., 2010), Pat1 (Pilkington & Parker, 2008), and Lsm4 (Ingelfinger et al., 2002).

All of these proteins possess multi-domains that mediate the assembly process via self-

interaction properties and interacting with other proteins. Three separate domains of Edc3

mediate the self-interaction and its association with other proteins including Dcp1 and the

RNA helicase Dhh1/RCK leading to an enhanced PB assembly (Decker et al., 2007; Kulkarni

et al., 2010). The Pat1 C-terminal domain promotes PB assembly via its interaction with

DCP1, while the middle domain of Pat1 interacts with Lms1 (Pilkington & Parker, 2008). In

addition, the Q/N-rich prion-like domain of Lsm4p is required for P-body assembly (Decker

et al., 2007). Decker et al, 2007 suggested that two sub-complexes of a conserved core of

proteins associate directly with mRNA playing an important role in the assembly of PBs, the

first complex consists of Dcp1p, Dcp2p, Dhh1p, and Edc3p and the second consists of Pat1p,

Lsm1-7p, and Xrn1p. The interaction between these two complexes via Edc3p YjeF-N

domain and/or the Lsm4p Q/N-rich prion-like domain is proposed to mediate PB formation

(Decker et al., 2007). Moreover, the YjeF-N domain is conserved amongst higher eukaryotes

promoting PB assembly (Ling et al., 2008). In metazoan, Lsm4 Q/N-rich prionlike domain is

not conserved and instead is replaced by Arg-Gly-Gly (RGG) involved in the SG assembly

process (Tourriere et al., 2003). Other proteins containing a Q/N region have been identified

as metazoan PB components including GW182 and Hedls/Ge-1 protein (Decker et al., 2007;

Eulalio et al., 2007b; Liu et al., 2005; Yu et al., 2005). In addition to GW182, other proteins

including RAP55 (also known as Lsm14) and Ge-1 (also known as Hedls or RCD-8) are

required for PB assembly in human cells, and depletion of these proteins leads to the

disassembly of PBs (Eulalio et al., 2007b; Yang et al., 2006b; Yu et al., 2005). In

mammalian cells, PB formation is greatly reduced by the depletion of GW182 and Ge-1,

which have no orthologous in yeast (Eulalio et al., 2007a), as well as Lsm1, RCK/p54 (Dhh1)

25

(Chu & Rana, 2006; Coller & Parker, 2005), and eIF4E-T (Andrei et al., 2005; Ferraiuolo et

al., 2005). Moreover, the disassembly of PBs is associated with the inhibition of miRNA

pathway, and the depletion of proteins involved in miRNA process, such as Drosha and its

binding partner DGCR8, lead to the dissolution of PBs (Pauley et al., 2006). PBs are highly

mobile structures that move along microtubules and the link between the intracellular

microtubule network and formation of PBs has been reported (Aizer et al., 2008; Aizer &

Shav-Tal, 2008; Baguet et al., 2007; Sweet et al., 2007). Aizer et al., 2008 demonstrated that

disrupting microtubules using the drug benomyl caused aggregation of PB components in

yeast (Sweet et al., 2007), which suggested that the enhancement of PB formation may occur

by preventing PB components from being trafficked away by microtubules. The same

relationship between cytoskeleton and assembly of PBs is also reported in mammalian cells

(Aizer et al., 2008; Aizer & Shav-Tal, 2008). These studies suggested that the formation of

PBs is controlled under normal conditions and that microtubules are involved in this process

to restrict the size or number of PBs. Furthermore, the assembly of RNA granules is

controlled by different signalling pathways. For example, the PKR pathway plays an

important role in the regulation of PB assembly/disassembly in yeast by phosphorylating the

Pat1 protein (Shah et al., 2014).

1.4.2 P-bodies function

The localisation of proteins associated with processes such as mRNA decay,

surveillance, silencing and translation repression in PBs has encouraged many researchers to

investigate any possible role for these cytoplasmic foci in these mechanisms. However, the

role of PBs varies depending on the particular cell system and stress condition that induces

these cytoplasmic granules (Shah et al., 2013).

26

1.4.3 Role of P-bodies in mRNA degradation

Eukaryotic cells employ two pathways for degrading mRNAs, both of which start

with shortening of the 3′ poly (A) tail (deadenylation) mediated by the Pan2-Pan3 and Ccr4-

Not complexes. Following deadenylation, mRNA can either undergo degradation from 5′-3′

by Xrn1, the 5′-3′ mRNA decay pathway, or degradation by the exosome from 3′-5′ after

removal of the 5′ cap structure by decapping enzymes, the 3′-5′ decay pathway (Balagopal &

Parker, 2009; Chen & Shyu, 2011; Coller & Parker, 2004; Wahle & Winkler, 2013)(Figure

1.2). To date, the actual role of PBs in mRNA decay is not yet clear, but the presence of 5′-3′

mRNA decay pathway enzymes, such as Dhh1, Xrn1, Dcp1/2, Pat1, Edc3, Lsm1-7 and Ccr4-

Not, strongly supports that these structures are potential sites for mRNA degradation (Jain &

Parker, 2013a). Several observations provide evidence to support this idea. For instance, the

appearance of PBs is associated with the presence of mRNA, and blocking transcription using

Actinomycin D or blocking mRNA decay by preventing deadenylation both affect the

assembly of PBs (Andrei et al., 2005; Cougot et al., 2004; Teixeira et al., 2005). Moreover,

the recognition of mRNA decay intermediates in PBs also supports the role of PBs in mRNA

decay (Sheth & Parker, 2003). The exosome and its Ski co-factors have not been identified as

PB components and therefore PBs are not associated with 3′-5′ degradation of mRNA

(Brengues et al., 2005; She et al., 2004). Furthermore, the PB components Pat1 and Lsm1-7

play a role in limiting exosome-mediated 3′-5′ mRNA degradation (He & Parker, 2001).

27

Figure 1.2 Eukaryotic mRNA decay pathways. The two general decay pathways in

eukaryotes include 3′ to 5′ and 5′ to 3′ degradation, both of which start with the

deadenylation of the 3′ poly (A) tail of the mRNA by the deadenylase

Ccr4p/Pop2p/Not complex. The deadenylation is followed by either 3′ to 5′

degradation by the exosome or by decapping by the 5’ decapping, mediated by

Dcp1/Dcp2 and 5′ to 3′ degradation by the exonuclease Xrn1. This second pathway is

thought to occur in P-bodies.

28

1.4.4 Role of P-bodies in mRNA surveillance

Nonsense-mediated decay (NMD) is one of the quality control mechanisms employed

by eukaryotic cells to terminate the translation of aberrant mRNA (Baker & Parker, 2004).

The surveillance complex involved in NMD includes UPF1-3 proteins and NMD effectors

SMG1/ 5/6/7 that induce termination of premature translation (Behm-Ansmant & Izaurralde,

2006). This complex is involved in the recruitment of functional proteins that promote

mRNA decay such as Xrn1 and decapping enzymes in yeast (Cao & Parker, 2003). The link

between mRNA surveillance and PBs is not fully clear, but data suggest that SMG7 localises

to PBs and induces the aggregation of SMG5 and UPF1 in these foci (Fukuhara et al., 2005;

Sheth & Parker, 2006). UPF1 also promotes the accumulation of UPF2 and UPF3 in PBs of

yeast cells lacking Dcp1 (Eulalio et al., 2007a). The accumulation of NMD proteins in PBs in

yeast may reflect the possible role for these RNA foci in mRNA surveillance (Eulalio et al.,

2007a). By contrast, Stalder and Mühlemann, 2009 have reported that the NMD process in

mammalian cells does not require PBs (Stalder & Muhlemann, 2009). PBs have also been

linked to [AU-rich element (ARE)-mediated mRNA decay] AMD, a decay process involving

ARE-containing mRNAs. TTP/BRF proteins may also mediate this degradation process (Lai

et al., 2000; Stoecklin et al., 2002). Depletion or knockdown of general decay pathway

proteins, Xrn1, Lsm1, Dcp1 or 4E-T all lead to inhibition of AMD and accumulation of ARE-

containing mRNAs and TTP/BRF proteins in PBs (Ferraiuolo et al., 2005; Franks & Lykke-

Andersen, 2007; Stoecklin et al., 2006).

1.4.5 Role of P-bodies in gene silencing

Small RNAs (~22 nt), including small interfering RNAs (siRNAs) and microRNAs

(miRNAs), involved in gene silencing, employ different effector mechanisms to silence their

29

target genes (Liu et al., 2005). This process is involved in post-transcriptional gene

expression regulation in higher eukaryotes (van Wolfswinkel & Ketting, 2010). The role of

PBs in RNA-mediated gene silencing is still a controversial issue (Leung & Sharp, 2013).

The localisation of specific RNA-induced silencing complexes (RISC) proteins such as

Argonaute proteins to PBs supports a role for PBs in RNA silencing (Liu et al., 2005).

Studies have demonstrated that Argonaute proteins also interact with other PB components

such as GW182, Dcp1, Dcp2 and RCK/p54, suggesting the presence of these proteins as a

complex in PBs (Eulalio et al., 2007a; Jakymiw et al., 2005; Liu et al., 2005). In human cells,

the depletion of GW182 or of the Dcp1-Dcp2 complex both impair miRNA function (Liu et

al., 2005). However, the depletion of other PB components such as Lsm1 or RCK/p54,

leading to loss of PBs has no effect on siRNA function (Chu & Rana, 2006; Jagannath &

Wood, 2009). Disruption of PB formation by trapping the mRNA in polysomes also did not

correlate with siRNA function (Chu & Rana, 2006; Franks & Lykke-Andersen, 2007).

1.4.6 Other functions of P-bodies

PBs are found to be important for the long-term survival of the cell and are present in

stationary phase in a process mediated by the PKA signalling pathway (Ramachandran et al.,

2011; Shah et al., 2013). PBs have also been proposed to be involved in the immunity

disorder (Bloch et al., 2013). A link between PB formation and translational repression or

regulation (Coller & Parker, 2005), and mRNA storage (Brengues et al., 2005; Bhattacharyya

et al., 2006) during viral infection has also been extensively reported (Tsai & Lloyd, 2014)

(see later section).

30

1.5 Interactions between stress granules and P-bodies

Despite differences between SGs and PBs in their composition (SGs contains 40S

ribosomal subunits and associated translation initiation factors such as eIF3 but these are not

present in PBs), size (PBs 100-300 nm and SG 0.2-5 μm), and mechanism of assembly and

function, a dynamic link between these cytoplasmic granules has been reported (Anderson &

Kedersha, 2006; 2009a; Kedersha et al., 2005; Wilczynska et al., 2005). SGs and PBs can

share certain proteins (such as eIF4E, RCK, TIA-1, TIAR, TTP, Xrn1 and Argonaute

proteins), making a clear distinction between them difficult, and they also contain specific

translationally repressed mRNA (Anderson & Kedersha, 2008; Buchan et al., 2008; Kedersha

et al., 2005; Mokas et al., 2009) (Figure 1.3). Although, direct evidence of mRNA transit

between these granules is still lacking, some studies have reported that the overexpression of

certain proteins such as TTP or BRF1 (Kedersha et al., 2005), and CPEB1 (Wilczynska et al.,

2005), mediated mRNA transit between these two cytoplasmic granules within the same cell.

This may reflect the presence of the same reporter mRNA in these foci, and these structures

would house mRNPs representing different stages of the mRNA life cycle rather than

different types of transcripts (Anderson & Kedersha, 2009a). Both granules form in the

cytoplasm of all eukaryotic somatic cells in response to several types of stress and their

formation is strongly associated with disassembly of polysomes (Hofmann et al., 2012;

Kedersha et al., 2005). Buchan et al., 2008 demonstrated that the SGs appear next to PBs in

S. cerevisiae (Buchan et al., 2008). In mammalian cells, the docking of these granules

together was also detected using live cell imaging experiments (Kedersha et al., 2005).

Furthermore, the live cell imaging also revealed an association and fusion events between

PBs and SGs (Kedersha et al., 2005; Wilczynska et al., 2005). According to these findings,

these cytoplasmic granules can transiently interact together (Anderson & Kedersha, 2009a).

31

Figure 1.3 Components of P-bodies and stress granules. The proteins markers found in

stress granules only (red), P-bodies only (yellow) and in both are displayed.

32

Because of the reverse correlation between disassembly of polysomes and assembly

of SGs or PBs and the dynamic link between these granules, as well as the ability of mRNA

to re-enter translation following relief from stress (Anderson & Kedersha, 2006;

Bhattacharyya et al., 2006; Brengues et al., 2005), several studies have suggested that the

eukaryotic mRNAs are in dynamic equilibrium between a translated pool (polysomes) and

non-translated pool either stored in SGs or degraded in PBs (Anderson & Kedersha, 2009a;

b; Balagopal & Parker, 2009; Brengues et al., 2005; Decker & Parker, 2012; Layana et al.,

2012; Lloyd, 2012) (Figure 1.4). Thus the mRNA life cycle between these subcellular

structures can affect the rate of mRNA translation and degradation (Beckham & Parker,

2008). Recently, Simpson et al., 2014 investigated the re-distribution of specific mRNA to

PBs after stress in live yeast cells using the mTAG system and reported that the movement of

specific endogenous mRNAs is biphasic, with some mRNAs present in RNA granules early

after stress, while other mRNAs localise much later in the stress period and these events were

associated with the recruitment of specific PB proteins such as eIF4E and Bfr1p (Simpson et

al., 2014). Distinguishing which mRNA species are present in SGs or PBs remains a matter

of debate (Thomas et al., 2011; Tsai & Lloyd, 2014). Available data reported the exclusion of

certain stress-active transcripts such as heat shock protein mRNA from SGs during heat stress

(Anderson & Kedersha, 2002; Nover et al., 1989). Other studies have confirmed the presence

of certain mRNA transcripts in SGs (Mazroui et al., 2007; Stohr et al., 2006). Many factors

may be involved in determining which mRNA transcripts will be targeted to RNA granules,

for example: the presence of proteins that harbour U-rich RNA such as TIA-1 (Buchan &

Parker, 2009; Cassola et al., 2007), which bind specific mRNA targeting them to SGs. More

recently, Lui et al., 2014 using live imaging of yeast cells, showed the localisation of specific

glycolytic mRNAs to cytoplasmic granules in unstressed growing cells. Following stress,

these granules coalesce and serve as a platform for PB assembly through the recruitment of

33

mRNA decay components. Interestingly, these mRNAs are translated within these

cytoplasmic granules under non-stress conditions and not associated with the mRNA decay

machinery. This suggests that the storage or decay of some mRNAs is driven by the pre-

existence of mRNP granules containing translationally active mRNAs (Lui et al., 2014).

Unlike PBs and SGs which harbour repressed mRNAs, the mRNA granules of unstressed

cells were unaffected by cycloheximide treatment which traps mRNAs in polysomes,

indicating that the ribosome may continue translation elongation in these granules (Lui et al.,

2014).

34

Figure 1.4 Proposed model for the role of cytoplasmic RNA granules in mRNA fate.

The mRNA is in dynamic equilibrium between a translated state (polysome) and un-

translated state (SGs and PBs). Under normal condition, the mRNA associates with

polysomes and is translated into polypeptides. Following stress, mRNAs can be stored in

cytoplasmic granules (SGs or PBs) to stall their translation. mRNAs can exit the SGs or

PBs to re-enter translation upon stress recovery or be further targeted to PBs for

degradation.

35

1.6 Stress granules, P-bodies, and viral life cycles

Viruses depend on the host cell for their replication cycle as they lack the

macromolecular machinery and small molecule precursors required to complete their

replication cycle (Birch et al., 2012). Therefore a variety of mechanisms are employed by the

infected host cells to cope with and restrict viral infection (Montero & Trujillo-Alonso, 2011;

Pattnaik & Dinh, 2013; Reineke & Lloyd, 2013; Tsai & Lloyd, 2014). The formation of SGs

and PBs in response to stress generated by viral infection was recently described as a novel

cellular mechanism to restrict viral infection (Beckham & Parker, 2008), silencing translation

thereby giving the cells more time to decide their fate via activation of other antiviral

signalling pathways as well as conserving their energy while accumulating proteins that are

required for adaptation (Pichon et al., 2012; Simpson & Ashe, 2012; Tsai & Lloyd, 2014). In

parallel, viruses have evolved different strategies to counteract, tolerate, or even take

advantage of these antiviral responses via the modulation of SG and PB formation during

infection (Beckham & Parker, 2008; Montero & Trujillo-Alonso, 2011; Schutz & Sarnow,

2007). Viruses can affect RNA granule formation either by disrupting the assembly of RNA

granules or utilizing RNA granule proteins for viral propagation (Reineke & Lloyd, 2013).

The understanding of the interplay between PBs, SGs and viral life cycle is still recent and

developing (Beckham & Parker, 2008; Lloyd, 2013; Reineke & Lloyd, 2013).

A large body of work supports this relationship through the identification of critical

proteins within PBs or SGs that promote or restrict viral infection, or by the demonstration

that some viral RNAs and proteins, as well as host antiviral defence proteins, accumulate in

PBs and/ or SGs (Beckham & Parker, 2008; Jain & Parker, 2013a; Kedersha & Anderson,

2007; 2009; Lloyd, 2013; Reineke & Lloyd, 2013; White & Lloyd, 2012). The sequestration

of mRNA in a translationally silenced state in both SGs and PBs affects viral mRNA fate;

therefore viruses have evolved a variety of mechanisms to avoid this fate (Lloyd, 2012). The

36

broad range of virus-RNA granule interactions is largely dependent on the viral system and

will be further discussed within the next sections (Pattnaik & Dinh, 2013; Tsai & Lloyd,

2014).

1.6.1 Viral interaction with stress granules

Viruses interact with SGs via a variety of strategies which may generally be of two

types: either a virus-mediated blockade of SG formation or viral replication that occurs

despite the presence of SGs because the virus is able to tolerate or exploit the SG response

(Montero & Trujillo-Alonso, 2011; White & Lloyd, 2012) (Figure 1.5).

The inhibition of SGs formation by viruses can occur throughout infection or in the

mid-phase of infection depending on the virus. In both cases, the induction of SGs usually

happens early in infection following the activation of eIF2 kinases by virus recognition

(Montero & Trujillo-Alonso, 2011). Later, the inhibition of SGs formation occurs either by

cleavage/modification of SG components or by sequestering key SG-nucleating components

that drive SG assembly (Lloyd, 2012; Tsai & Lloyd, 2014). Additionally, the manipulations

of PKR or in some cases PERK also contribute to the inhibition of stress granules formation

(Lloyd, 2013). Interestingly, some viruses benefit from the SG-mediated antiviral response by

recruiting specific SG components to their replication complex to enhance their replication

cycle (Emara & Brinton, 2007; Li et al., 2002; Panas et al., 2012; Yi et al., 2011). Moreover,

a complex oscillation between SG assembly and disassembly has also been reported during

certain viral infection (Ruggieri et al., 2012).

37

Figure 1.5 Mechanisms of stress granules disruption by viruses. Viruses have evolved

several mechanisms to disrupt or prevent SGs assembly. (A) HCV, flavivirus, JUNV,

alphavirus and VSV can all induce sequestration of different SG components to their

replication complex (RC). (B) The picornavirus proteases can cleave nucleating-SG proteins

leading to inhibit SG assembly. (C) Viral proteins can interact with SGs components or SG-

inducing pathways to prevent the formation of SGs. (D) The re-localisation of SG proteins

from cytoplasmic foci to nucleus can also affect the formation of SGs as in the case of IAV

and rotavirus.

38

1.6.2 Inhibition of Stress granules by viral proteinase

Three picornaviruses including poliovirus (PV), coxsackievirus B3 (CVB3) and

encephalomyocarditis virus (EMCV) have been found to induce stress granule formation

early in infection (2-3 h.p.i), independently of eIF2 phosphorylation (Fung et al., 2013; Ng et

al., 2013; White et al., 2007). Later, the disassembly of these granules during the middle

phase of the viral replicative cycle (3-8 h.p.i) occurs via the cleavage of G3BP by the viral 3C

proteinase at the residue glutamine 326, thereby preventing the localisation of mRNA and

associated factors such as the initiation factors eIF4G and PABP and 40S ribosomal subunits

to cytoplasmic foci (White et al., 2007). In contrast, the aggregation of TIA-1 still occurs at

late times post-infection in PV-infected cells forming unconventional SGs that lack G3BP,

eIF4G and PABP (White et al., 2007; White & Lloyd, 2011). The treatment of PV-infected

cells with actinomycin D, an inhibitor of cellular transcription has a negative impact on SG

formation (Piotrowska et al., 2010), suggesting that cellular transcription may be important

for SG assembly during PV infection. More recently, Wu et al., 2014 have demonstrated that

the viral proteinase 2Apro is necessary to trigger SG formation early in infection (3 h.p.i) by

both CVB3 and enterovirus 71 (EV71), suggesting that other picornaviruses might use the

same mechanism to initiate SG responses (Wu et al., 2014). While the exact mechanism of

2Apro-mediated SG formation remains unclear, 2Apro proteinase activity is critical to initiate

SGs formation (Wu et al., 2014). It was proposed that this may be via the ability of 2Apro to

re-localize SRp20, a nuclear-cytoplasmic shuttling splicing factor, from the nucleus to the

cytoplasmic granules (Fitzgerald et al., 2013), or through the cleavage of eIF4G by 2Apro (Wu

et al., 2014). Other studies have also provided evidence for the absence of G3BP1 and eIF4G

from SGs at late stages of CVB3 and EV71 infections (White et al., 2007; Wu et al., 2014).

39

1.6.3 Inhibition of stress granule formation by modulation of eIF2 phosphorylation

Responding to viral entry, mammalian orthoreovirus (MRV), a Reoviridae, induces

SG formation early during infection (6hpi) and this correlates with an increase in eIF2

phosphorylation and reduced cellular translation (Qin et al., 2011). These events may

promote viral uncoating which depends on eIF2 phosphorylation and expression of ATF4, a

transcriptional factor that enhances viral replication when cellular mRNAs localise to SGs

(Smith et al., 2006b). Interestingly, at this stage SGs contain viral core particles but the role

of this is unclear (Qin et al., 2009). Associated with an increase of viral translation, the

disassembly of SGs is observed late in infection (24 hpi) as the replication cycle progresses

independently from eIF2α phosphorylation (Qin et al., 2011). Moreover, the knockdown of

PKR or other eIF2α kinases has no effect on SG formation, indicating that the induction of

SGs during MRV infection may occur by involving two or more eIF2α kinases and through

the contribution of viral proteins (Qin et al., 2009). These findings clearly highlight that

MRV regulates assembly/disassembly of SGs as the replication cycle progresses. The

induction of eIF2α-dependent SGs early during infection and their blockage late in infection

was also shown in mouse embryo fibroblasts (MEF) during Semliki Forest Virus (SFV)

infection (McInerney et al., 2005).

Recently, Ruggieri et al., 2012 developed a live-cell imaging approach to demonstrate

that HCV utilises specific mechanisms to control SG assembly/disassembly throughout the

infection cycle via the oscillation of SG formation, preventing prolonged translation

repression and promoting viral replication. This mechanism depends on DNA damage–

inducible protein 34 (GADD34), the cofactor of phosphatase1 that dephosphorylates eIF2α

and PKR, regulating the phosphorylation status of eIF2α (Ruggieri et al., 2012). Junín virus

(JUNV) is another example of a virus that mediates blockage of SG formation. JUNV

40

infection does not induce SG formation in Vero cells, and the induction of SGs is impaired in

infected cells exposed to sodium arsenite-mediated stress (Linero et al., 2011). Studies in

transfected cells revealed that the inhibition of SGs during JUNV infection is mediated by

two viral proteins, nucleoprotein or N protein and glycoprotein precursor (GPC) which affect

eIF2α phosphorylation (Linero et al., 2011). By contrast, the assembly of SGs was observed

in JUNV- infected cells treated with the eIF4A inhibitor, hippuristanol, which induces SG

formation independently from eIF2α phosphorylation (Linero et al., 2011).

1.6.4 Inhibition of stress granule formation by sequestering or co-opting their

components

The sequestration of the SG protein G3BP1 through its interaction with both NS5B

protein of hepatitis C virus (HCV) and the 5′ end of the HCV minus-strand RNA, late in

infection (48 hpi), causes inhibition of SGs induced earlier in infection via PKR activation

(Yi et al., 2011). This study also demonstrated that G3BP1 co-localizes with HCV replication

complexes (RCs) in cells expressing an HCV replicon. HCV replication was dramatically

reduced by siRNA-mediated knockdown of G3BP1, indicating that RC-associated G3BP1

may be co-opted as a functional protein for viral replication.

The sequestration of G3BP1 to RC during SFV infection via its interaction with the

C-terminal domain of the viral non-structural protein 3 (nsP3) was also found to reduce the

formation of SGs early in infection and inhibit their formation late during infection (Panas et

al., 2012). Another example is human T-cell leukaemia virus type 1 (HTLV-1), a

Retroviridae member which uses a different strategy to block SG formation involving

sequestration of another protein critical for assembly of SGs. This strategy is mediated by

Tax protein which is found in the nucleus under normal conditions but shuttles to the

41

cytoplasm in response to different types of stress (Legros et al., 2011). Within the cytoplasm,

Tax associates with histone deacetylase 6 (HDAC6), a protein involved in SGs assembly

(Kwon et al., 2007), thereby impairing SGs formation (Legros et al., 2011). Cricket paralysis

virus (CrPV), a member of the Dicistroviridae family also blocks SGs formation by

preventing Rox 8 and Rin, the Drosophila homologs of TIA-1 and G3BP1, respectively, from

localising in infected Drosophila S2 cells, even in the presence of stress factors such as heat

shock, arsenite or pateamine A (Khong & Jan, 2011). Other flaviviruses, such as West Nile

virus (WNV) and dengue virus (DV), also have RC colocalizing with TIA-1 and TIAR

proteins via an interaction with the 3′ end of the WNV viral genome thus impairing SGs

assembly (Emara & Brinton, 2007; Li et al., 2002). These viruses take advantage of this

antiviral response, because the lack of these proteins, for example TIAR-depletion in MEF

cells, impairs viral replication (Li et al., 2002). In another flavivirus, Japanese encephalitis

virus (JEV), the interaction between the core structural protein and Caprin-1 (a SGs

component) mediates the inhibition of SGs formation via the recruitment of several SG-

associated proteins, including Caprin-1, G3BP1 and USP10, thereby promoting viral

propagation (Katoh et al., 2013).

Rotavirus has evolved a mechanism to block SGs despite eIF2 phosphorylation

during the early stage of infection (Rojas et al., 2010). Montero et al., 2008 suggested that

this overcoming of eIF2 phosphorylation-mediated translation arrest may be due to the

relocation of PABP (a component of SGs) from the cytoplasm to the nucleus thereby

preventing the formation of SGs (Montero et al., 2008).

In the case of human immunodeficiency virus 1 (HIV-1) infection, Staufen1 is

recruited to RNPs containing genomic RNA via its interaction with Gag inhibiting SGs

formation (Abrahamyan et al., 2010). The mechanism by which HIV-1 blocks SGs assembly

42

also involves the interaction of viral Gag protein with eukaryotic elongation factor 2 (eEF2)

and this process occurs irrespective of eIF2α phosphorylation or overexpression of G3BP1 or

TIAR, which generally trigger SGs formation (Valiente-Echeverria et al., 2014). By contrast

with HIV-1, HIV-2 expression induces stress granule assembly. Moreover, HIV-2 genomic

RNA associates with TIAR to form a TIAR-HIV-2 ribonucleoprotein (TH2RNP), which is

diffused in the cytoplasm or accumulated in SGs in the absence of active translation (Soto-

Rifo et al., 2014). Cycloheximide or puromycin treatment lead to either complete localization

of the TH2RNP complex diffused throughout the cytoplasm, or the incorporation of this

complex in enlarged SGs indicating the TH2RNP complex is in equilibrium between

polysomes and SGs. Nevertheless, the formation of TH2RNP complex does not require

Gag/gRNA binding, but the level of the Gag polyprotein could regulate translation and the

cytoplasmic localization of this complex. A low concentration of Gag promotes the

translation of gRNA and diffused localisation of the gRNA and TIAR in the cytoplasm,

whereas a high level of Gag inhibits the translation of gRNA leading to accumulation of

gRNA/TIAR in SGs. The ability of the gRNA and Gag to accumulate in SGs may reflect

their role as sites where the switch from viral translation to packaging occurs (Soto-Rifo et

al., 2014).

Theiler’s murine encephalomyelitis virus (TMEV), a picornavirus, inhibits SGs

assembly induced by sodium arsenite or thapsigargin treatment in a process mediated by the

leader (L) protein (Borghese & Michiels, 2011). TMEV viral L protein inhibits the host

immune response by disrupting IFN and chemokine production thereby enhancing viral

persistence (van Pesch et al., 2001). Interestingly, the L protein prevents SG formation

independently from its inhibition of interferon production. This raised the possibility that the

inhibition of SGs formation is due to the role of the L protein in altering nucleocytoplasmic

trafficking factors involved in SG assembly (Borghese & Michiels, 2011). NS1 protein-

43

mediated PKR inhibition is essential to block SG formation at a late stage of influenza A

virus (IAV) infection (Khaperskyy et al., 2012). Khaperskyy et al., 2014 demonstrated that

two other proteins can inhibit SGs formation during IAV infection and that the nucleoprotein

(NP) and host-shutoff protein polymerase-acidic protein-X (PA-X) inhibit SGs formation

independently from eIF2α phosphorylation. NP and PA-X action are concomitant with a

dramatic decrease and increase of PABP in the cytoplasm and nucleus respectively

(Khaperskyy et al., 2014). Furthermore, the interaction between NS1 and RNA-associated

protein 55 (RAP55) also impairs SGs formation (Mok et al., 2012).

1.6.5 Viral replication mechanism that benefits from stress granule formation

Some viruses can replicate and cope with the presence of SGs during infection. For

example, despite some effect, the mouse hepatitis coronavirus (MHE) replicative cycle

continues during infection while SGs are stimulated through eIF2 phosphorylation (Raaben

et al., 2007). The replication of respiratory syncytial virus (RSV) is also completed despite

the presence of SGs induced via PKR-mediated mechanisms (Lindquist et al., 2011).

In the case of negative-stranded RNA viruses, Vesicular stomatitis virus (VSV)

infection induces unconventional SGs because these granules do not contain some of the

bona fide SG markers, such as eIF3 or eIF4A, or the processing body (PB) markers, Dcp1a

and so-called SG-like structures which contain poly-C binding protein (PCBP2), TIA1 and

TIAR. Interestingly, VSV SG-like structures contain the viral replicative proteins (VSV N

protein) and viral RNA. Moreover, viral replication and protein synthesis as well as the

presence of TIA-1 are necessary for forming the SG-like structures during VSV infection

(Dinh et al., 2013). In this study, the formation of SG-like structures had no significant effect

44

on viral protein synthesis despite increased levels of phosphorylated eIF2 at different times

post-infection.

All the examples mentioned above are RNA viruses; few studies have investigated the

relationship between SGs formation and DNA viruses (Esclatine et al., 2004; Isler et al.,

2005). For example, herpes simplex virus-1 (HSV-1), SGs are not observed during infection

(Esclatine et al., 2004). However, the induction of SGs using sodium arsenite, a classical

inducer of oxidative stress is also inhibited in HSV-2 infected cells, whereas the SGs

containing G3BP1 and PABP but not TIA-1 were observed in infected cells that treated with

Pat A (Finnen et al., 2012). This may be due to an interaction of virion host shutoff (Vhs)

protein (HSV virion component) with a key component of SGs such as TTP (Dauber et al.,

2011; Esclatine et al., 2004; Finnen et al., 2014). The infection with vhs-deficient mutants of

HSV-1 or -2 induces SGs formation late in infection, these granules contain core SG

components such as TIA-1, TIAR and G3BP1 as well as the viral immediate early protein

ICP27 and the viral serine/threonine kinase Us3 (Esclatine et al., 2004; Finnen et al., 2014).

45

1.7 Virus/P-bodies interaction

Compared with SGs, there are only a few studies describing the relationship between

PBs and viruses, most of which are related to positive-stranded RNA viruses (Pattnaik &

Dinh, 2013; Tsai & Lloyd, 2014). In addition to the role of PBs in suppressing certain viral

life cycles, other mechanisms for the disruption of PBs during viral infection or viral co-

opting of PBs components have been described (Lloyd, 2013) (Figure 1.6).

In addition to 3CPro cleaving G3BP1 during PV infection, Dougherty et al, 2011 also

proposed a role for this enzyme in the rapid disruption of PBs 2-4h after PV and CVB3

infection through the cleavage or degradation of key components of PBs such as Xrn1, Dcp1a

and Pan3, suggesting an inhibition of the de-adenylation which may be required for viral

replication (Dougherty et al., 2011). In addition, WNV and DV impair PB assembly at a late

stage of infection (24-36 hpi) and the reduction of PBs was shown to be concomitant with the

relocation of TIA-1 and decreased eIF2 phosphorylation level in cells exposed to oxidative

stress via unidentified mechanisms (Emara & Brinton, 2007). Later, Chahar et al., 2013

reported that WNV infection causes the recruitment of many PB components including

Lsm1, GW182, DDX3, DDX6 and Xrn1 that localise with viral non-structural protein 3

(NS3) at viral replication sites. Some of these proteins have been demonstrated to be positive

regulators of viral replication as their siRNA- mediated knockdown impairs viral replication

(Chahar et al., 2013). The progressive decrease in PBs was also observed at a late stage

during HCV infection. DDX6 or G3BP1 were found to colocalise with HCV core 48 h after

infection (Ariumi et al., 2011), indicating that the HCV hijacks the P-body and the stress

granule components to regulate HCV RNA replication and translation. Supporting this,

knockdown of DDX3 reduced HCV replication. Other PB constituents including Ago2,

Lsm1, PatL1, Ge1, and miR122 are required for HCV replication as their knockdown had a

46

negative impact on viral replication (Ariumi et al., 2011; Berezhna et al., 2011; Pager et al.,

2013; Thibault et al., 2013).

Recently, using hepatocytes from human livers of patients infected with different

genotypes of HCV, Perez-Vilano et al., 2015 were also able to demonstrate disruption of P-

body formation in vivo (Perez-Vilaro et al., 2014). They also demonstrated that this inhibition

was relieved upon HCV elimination using antiviral therapy. Interestingly, they also provided

evidence for a high heterogeneity in the composition and shape of PBs formed in uninfected

hepatocytes. This clearly demonstrates a link between PBs formation and a key pathogenic

condition.

Other examples of viruses that co-opt components of PBs to enhance their replication

are HIV and MLV, as the replication of both viruses is affected by the depletion of Mov10, a

PBs protein (Burdick et al., 2010; Furtak et al., 2010). Moreover, HIV virion assembly is

reduced by knockdown of DDX6 and Ago2 which both localise with HIV Gag protein at PBs

assembly sites (Reed et al., 2012). Among negative-sense RNA viruses, NS1 viral protein

mediates the disruption of PBs during influenza virus infection via its interaction with Rap55,

preventing the sequestration of the viral NP and NP-associated RNAs in PBs thereby

promoting viral replication (Mok et al., 2012). The Hantavirus nucleocapsid protein binds to

the 5′ cap of cellular mRNAs and protects the 5′ extremity from decapping and degradation

during accumulation in PBs, and is involved in initiation of viral transcription (Mir et al.,

2008).

Less is known about the relationship between PBs and DNA viruses. However, the

adenovirus replication cycle causes PB reduction by altering PB components through a

specific interaction between viral proteins and DDX6. This association causes the

redistribution of several PB constituents to aggresomes (Greer et al., 2011).

47

Figure 1.6 Mechanisms of P-bodies disruption by viruses. Viruses evolved many

mechanisms to disrupt or prevent PBs. (A) HCV, flavivirus, and adenovirus can induce the

sequestration of different PBs components to their replication complex (HCV and flavivirus)

or to agressome (adenovirus), disrupting PBs assembly. (B) Viral protease from picornavirus

cleavage an important nucleating-PB proteins including Dcp1, Xrn1 and Pan3 disrupting PBs

assembly. (C) Interactions between specific viral proteins and PB components also prevent

formation of PBs.

48

1.8 Caliciviruses

The Caliciviridae were adopted as a family of viruses in the third report of the

International Committee on Taxonomy of Viruses (ICTV) in 1979, and before this

caliciviruses were considered as part of the Picornaviridae family (Green et al., 2000).

According to the last 2011 report of ICTV, the caliciviruses can be classified into five genera

(shown in Table 1.3) based on their genomic organisation and sequence similarities. These

genera are Lagovirus, Vesivirus, Norovirus, Sapovirus and Nebovirus (Green, 2013;

International Committee on Taxonomy of Viruses, http://ictvonline.org). Five additional

genera have been proposed including "Salovirus" (Mikalsen et al., 2014), "Bavovirus"(Wolf

et al., 2012), "Nacovirus" (Day et al., 2010; Wolf et al., 2012), "Recovirus" (Farkas et al.,

2014; Farkas et al., 2008) and "Valovirus" (L'Homme et al., 2009), and several Caliciviridae

remain unclassified (including 18 species; http://www.caliciviridae.com).

Caliciviruses are small, non-enveloped, positive-sense single-stranded RNA viruses.

Their genomes range between 7.3 to 8.5 kb in length, containing 2 to 4 open reading frames

(ORFs), and are surrounded by a small icosahedral capsid of 27 to 40 nm in diameter

(Rohayem et al., 2010; Thackray et al., 2007). The name calicivirus originates from the Latin

word for cup (“calyx”) because of the cup-shaped indentations observed on the virion using

electron microscopy (Thiel & Konig, 1999) (Figure 1.7)

The Caliciviridae family includes viruses that cause gastrointestinal disease in human

hosts and a broad spectrum of diseases in a wide variety of animal hosts (Glass et al., 2000b;

Rohayem et al., 2010) leading to significant economic impact both to the UK National Health

Service (NHS) and businesses in the case of human disease, and to the agricultural industry

and farming for animals.

49

Table 1.3 Taxonomy of the Caliciviridae family (ICTV, 2011); Caliciviruses taxonomy

browser, 2014)

Genus Species Viruses

Vesivirus

(2 Species - 46 complete

genomes)

Feline calicivirus

(711 strains - 32 complete genomes) Feline calicivirus serotypes

Vesicular exanthema of swine virus

(99 strains - 6 complete genomes)

Vesicular exanthema of swine virus serotypes, bovine calicivirus,

cetacean calicivirus, primate calicivirus, reptile calicivirus, San Miguel

sea lion virus serotypes

Sapovirus

(1 Species - 40 complete

genomes)

Sapporo virus

(2162 strains - 38 complete genomes)

Porcine enteric calicivirus, Sapporo virus, Houston virus, Manchester

virus

Norovirus

(1 Species - 688 complete

genomes)

Norwalk virus

(17729 strains - 624 complete

genomes)

Murine norovirus, Lordsdale norovirus, Norwalk virus, Snow

Mountain norovirus, Southampton norovirus

Lagovirus

(2 Species - 41 complete

genomes)

European brown hare syndrome virus

(82 strains - 4 complete genomes) European brown hare syndrome virus serotypes

Rabbit haemorrhagic disease virus

(547 strains - 35 complete genomes) Rabbit haemorrhagic disease virus serotypes

Nebovirus

(1 Species - 5 complete

genomes)

Newbury-1 virus

(15 strains - 4 complete genomes) Newbury-1 virus, Nebraska virus

Unclassified Caliciviridae

(18 Species - 20 complete

genomes)

(18 Species - 20 complete genomes)

Calicivirus strain HP167-KOR, Calicivirus strain MA120-KOR,

Calicivirus strain MA129-KOR, Calicivirus strain MA137-KOR,

Calicivirus strain MA16-KOR, Calicivirus strain MA64-KOR,

Calicivirus strain MA76-KOR, Calicivirus strain MA176-KOR,

Calicivirus strain MA216-KOR, Calicivirus strain MA236-KOR,

Calicivirus strain MA360-KOR, Calicivirus strain MA56-KOR

Calicivirus strain MA91-KOR, Calicivirus strain MA95-KOR,

Calicivirus strain MA99-KOR, Calicivirus strain SA181-KOR,

Calicivirus strain SA68-KOR, Calicivirus strain MA129-KOR,

Recombinant virus 6918 VP60-T2

50

Figure 1.7 Crystal structures of Norwalk Virus, MNV-1 and FCV by cryo-electron

microscopy from (Bhella et al., 2008; Katpally et al., 2008; Prasad et al., 1999). (A)

Structure of HuNoV (Norwalk Virus); (B) Structure of MNV-1; (C) Structure of FCV. The

cup-shaped indentations characteristics of caliciviruses are visible in green in A, blue in B,

and pink in C.

51

1.8.1 Lagoviruses (LaVs)

The genus Lagovirus consists of two species, Rabbit Haemorrhagic disease virus

(RHDV) and European Brown Hare Syndrome Virus (EBHSV) (ICTV, 2013). RHDV was

first detected in China in 1984 and associated with the death of 14 million domesticated

angora rabbits within nine months. The virus then rapidly spread worldwide via Italy to the

rest of Europe and is now present as an endemic agent in many countries (Abrantes et al.,

2012a; Abrantes et al., 2012b). All pathogenic strains are classified within a single serotype.

RHDV causes high mortality in both domestic and wild adult animals (Abrantes et al., 2012a;

McIntosh et al., 2007). The disease is characterised by acute necrotising hepatitis, but other

organs, such as lungs, heart, and kidney may also be affected (Abrantes et al., 2012b). The

transmission of RHDV is mainly via direct contact with infected rabbits through the oral

route and skin lesion (Merchan et al., 2011), or indirectly via contaminated food, bedding,

water, clothes and veterinary equipment (Cooke, 2002). Despite the similarities between

RHDV and EBHSV in clinical signs, pathological and histopathological alterations, mortality

rates, virion morphology and antigenicity, these viruses infect different species (Abrantes et

al., 2012b). EBHSV was first detected in Sweden in 1980 (Gavier-Widen & Morner, 1991),

and is the causative agent of European brown hare syndrome in Lepus europaeus and Lepus

timidus (Lopes et al., 2014). Subsequently, the virus was recognised in many European

countries (Paci et al., 2011) and now is considered endemic in all European countries (Chiari

et al., 2014). EBHSV is a highly contagious disease and transmission occurs via direct

contact or through the oral-fecal route (Frolich et al., 2007).

52

1.8.2 Vesiviruses (VeVs)

The Vesivirus genus is represented by two species, Vesicular exanthema of swine

virus (VESV), which consists of many serotypes and feline calicivirus (FCV) with a single

serotype. In addition, a number of vesiviruses remain unclassified (ICTV, 2013). VESV was

first observed in 1932 in California, USA, associated with vesicular disease in swine. From

1952 the disease spread through most USA states leading to massive economic losses (Smith

et al., 2006a). Later in 1972, related viruses were isolated from sea lions on San Miguel

Island, California and named as San Miguel Sea lion virus (SMSV), causing vesicular disease

when transferred to swine (Smith et al., 1973). This led to the suggestion that all cases of

VESV in USA between 1932 and 1959 were caused by marine origin serotypes (Reid et al.,

2007; Smith et al., 2006a). Importantly, Vesivirus of marine origins have been found to infect

humans (Smith et al., 2006a). In addition to the SMSV serotype, VESV species have several

serotypes infecting a wide range of hosts such as fish, reptiles, primates and others animals

(Smith et al., 2006a; Wellehan et al., 2010).

A second species of Vesivirus is feline calicivirus (FCV), a highly infectious pathogen

that infects domestic and wild cats, and is responsible for acute oral and upper respiratory

tract disease with a range of clinical signs which depend on the virulence of FCV strains

(Geissler et al., 1997; Radford et al., 2009). Some strains are associated with chronic

stomatitis and limping syndrome (Radford et al., 2009). The routes of transmission are nasal,

oral or conjunctival routes, and following infection, the replication occurs in the oropharynx.

The infection starts with a necrosis in epithelial cells which subsequently leads to ulcer

formation (Radford et al., 2009). FCV can cause pneumonia and lameness (Dawson et al.,

1994). Recently, highly virulent strains of FCV with high mortality (up to 67%) have been

recognised in the USA and Europe and cause FCV-associated virulent systemic disease

(FCV-VSD). They are responsible for outbreaks of severe and acute systemic disease which

53

differs from the more usual FCV disease (Geissler et al., 1997; Ossiboff et al., 2007;

Pesavento et al., 2004; Radford et al., 2007). These strains are characterised by a new range

of clinical signs including systemic inflammatory response syndrome, jaundice, ulcerative

dermatitis and oedema that can lead to death (Coyne et al., 2012; Pesavento et al., 2004;

Radford et al., 2009). Vaccination can prevent the disease but does not prevent the infection

(Coyne et al., 2006; Southerden & Gorrel, 2007). To explain this virulence, it was found that

the most variable region of FCV is located in the surface of the capsid protein, representing

the main target for the immune response (Radford et al., 2009). The evolutionary rate of this

region is up to 1.3x10-2 to 2.6x10-2 substitution per nucleotide per year, the highest

evolutionary rate reported compared with any virus (Coyne et al., 2012). In addition, FCV

displays resistance to common disinfectants and can persist in the environment for 1 month

(Radford et al., 2009). Importantly, FCV is used as a model for the study of the molecular

biology of human norovirus (Vashist et al., 2009).

1.8.3 Sapoviruses (SaVs)

The Sapovirus genus consists of a single species divided into five genogroups (GI-V),

and subdivided into a number of genotypes based on the capsid gene sequence (Hansman et

al., 2007; Oka et al., 2012). Saporoviruses are responsible for enteric diseases in human and

animal hosts (Hansman et al., 2007; L'Homme et al., 2010). To date, only genogroup GIII is

associated with animal disease especially in porcine species (Di Bartolo et al., 2014; Farkas

et al., 2004; Hansman et al., 2007; L'Homme et al., 2010; Pang et al., 2009). The other

genogroups (GI, GII, GIV and GV) have been reported as causative agents of human acute

gastroenteritis worldwide (Bucardo et al., 2014; Hansman et al., 2007; Nidaira et al., 2014;

Svraka et al., 2010; Usuku & Kumazaki, 2014; Wang et al., 2014). They can affect all age

54

groups but are more common in children (Dey et al., 2012; Hansman et al., 2007; Lee et al.,

2012; Sakai et al., 2001; Usuku & Kumazaki, 2014; Wu et al., 2008). Several reports have

also reported outbreaks of SaV in adults (Lee et al., 2012; Nidaira et al., 2014; Wang et al.,

2014). The first report of an SaV outbreak was by Chiba et al., 1977 using electron

microscopy of stool specimens from infants suffering from acute gastroenteritis in an

orphanage in Sapporo, Japan (Chiba et al., 1979). The genogroups GI/II have been reported

as the most frequent strains associated with outbreaks of gastroenteritis worldwide (Hansman

et al., 2007; Lee et al., 2012; Miyoshi et al., 2010; Nidaira et al., 2014; Ootsuka et al., 2009;

Usuku & Kumazaki, 2014; Wang et al., 2014).

Porcine SaV (PSaV) has been detected in symptomatic or diarrheic pigs (Collins et

al., 2009; Di Bartolo et al., 2014; L'Homme et al., 2009; Scheuer et al., 2013) and recently

numbers of novel animal SaV strains related to GIII and human strains have been identified

in mink and swine (Guo et al., 2001; Martella et al., 2008; Wang et al., 2005; Yin et al.,

2006). In addition, PSaV can be propagated in tissue culture, providing a good model for

understanding the molecular aspects of human SaV (Chang et al., 2002; Hosmillo et al.,

2014). More recently, SaV GI viruses have been detected in faecal samples from

chimpanzees living in close contact with humans in the Democratic Republic of Congo

(Mombo et al., 2014). Human SaV strains have been reported in food and water in areas

surrounding populations that have suffered from SaV outbreaks (Iizuka et al., 2010;

Kobayashi et al., 2012; Murray et al., 2013; Sano et al., 2011). Therefore, it is likely that the

transmission routes are faecal-oral as well as person-to-person (Sala et al., 2014). The close

genetic relationship between the SaV identified recently in both animals and humans, as well

as in food and water, may trigger further research to address the zoonotic origins of SaV

infections (Bank-Wolf et al., 2010).

55

1.8.4 Neboviruses (NeVs)

The most recent genus of the Caliciviridae family to be established, Nebovirus

contains a single species, Newbury-1 virus (Carstens, 2010). It comprises several strains such

as Newbury “agent”-1, first described in the UK in 1976 and Nebraska virus first detected in

Nebraska, USA in 1980 (Carstens, 2010; Oliver et al., 2006; Smiley et al., 2002). So far, all

strains described in this genus are bovine enteric viruses (Smiley et al., 2002) ICTV, 2013).

Recently, the frequent detection of an association between bovine caliciviruses such as

Nebovirus and Bovine norovirus (BNoV) with calf diarrhoea suggested that these viruses are

a significant cause of endemic diarrheal disease in calves (Cho et al., 2013; Kaplon et al.,

2011; Park et al., 2007). As the majority of caliciviruses, Nebovirus cannot be propagated in

cell culture (Carstens, 2010).

1.9 Noroviruses (NoVs)

NoVs are the most important causative agent of viral gastroenteritis outbreaks in

humans worldwide, but have also been detected in some animal hosts (Hall et al., 2011;

Ahmed et al., 2014; Green et al., 2000). This genus contains only one species, based on

phylogenetic analysis of their capsid gene (and recently of ORF1 sequence), which can be

divided into six genogroups (GI-GVI) containing several genotypes (Caddy et al., 2014;

Green et al., 2000; Kroneman et al., 2013; Zheng et al., 2006)(Figure 1.8). Animal NoVs

belong to genogroups GIII (bovine NoV), GV (murine NoV), GII (porcine NoV) and GIV

(lion NoV) (Caddy et al., 2014; Martella et al., 2007; Scipioni et al., 2008; Wang et al., 2005;

Zheng et al., 2006), whereas human NoVs fall in GI, II and IV (Atmar, 2010; Caddy et al.,

2014; Green et al., 2000; Zheng et al., 2006). Genetic relationships between human and

animal NoVs may reflect a potential zoonotic transmission (Scipioni et al., 2008).

56

Figure 1.8 Phylogenetic analysis of the capsid amino acid sequence of Norovirus

strains. Norovirus can be classified into six genogroups (GI-GVI) based on their capsid

gene and ORF1 sequence containing several genotypes as indicated. Genotype GII.4 has

seven different variants of genotypes as boxed. (Modified from (Glass et al., 2009;

Kroneman et al., 2013; Patel et al., 2009; Ramani et al., 2014).

57

1.9.1 Animal noroviruses

To date, animal noroviruses have been detected in cattle, pigs (Karst et al., 2003;

Oliver et al., 2003; Wang et al., 2005; Wolf et al., 2009), dogs and rats (Martella et al., 2007;

Pinto et al., 2012; Tse et al., 2012), lion cubs (Martella et al., 2007) and mice (Karst et al.,

2003), representing infection by bovine, porcine, canine, feline and murine noroviruses.

Diarrhoea is the common clinical sign reported in calves infected with NoV but is not

reported in infected pigs or mice (Dastjerdi et al., 1999; Hsu et al., 2007; Wang et al., 2007;

Wang et al., 2006)

1.9.2 Murine norovirus-1 (MNV-1)

MNV-1 was first discovered in 2003 in the USA and associated with lethal infection

of laboratory mice (Karst et al., 2003). Many studies have since then reported MNV

infections in laboratory mice in different countries (Hsu et al., 2005; Mahler & Kohl, 2009;

Ohsugi et al., 2013). Although MNV-1 can infect healthy mice it does not result in any

disease (Thackray et al., 2007). Mice lacking recombination-activation gene 2 (RAG2) and

activator of transcription 1 (STAT1), two factors involve in the immune responses, are very

sensitive to MNV-1, which can replicate within lymphoid tissues and display many clinical

signs leading to death (Hsu et al., 2006; Karst et al., 2003). This indicates that the innate

immune system components have a very strong effect on MNV-1 infection. To date, 15

distinct MNV strains have been isolated such as MNV-2, 3 and 4 (Hsu et al., 2006; Thackray

et al., 2007). These strains have different biological properties or pathogenic patterns but all

belong to one MNV serotype (Goto et al., 2009; Hsu et al., 2006; Thackray et al., 2007).

Transmission of MNV is mainly via the faecal-oral route especially through contaminated

bedding (Manuel et al., 2008). It has been demonstrated that MNV strains are able to

58

replicate and persist in different cells and tissues such as macrophages and dendritic cells

(Wobus et al., 2004) as well as the small intestine, caecum, mesenteric lymph nodes and the

spleen of both immune-competent and immune-deficient mice (Hsu et al., 2006; Thackray et

al., 2007). Other studies have reported that MNV circulates in wild rodents (Farkas et al.,

2012; Smith et al., 2012). MNV is the only norovirus that replicates in cultured cells, mainly

macrophage cell lines; MNV-1 can be propagated in RAW 264.7, BV-2 and TIB cell lines

(Cox et al., 2009). Because of these properties, and the genetic relationship to human NoVs,

MNV provides an excellent model to study the translation and replication mechanisms of

Human NoVs (Thorne & Goodfellow, 2014; Wobus et al., 2006).

1.9.3 Human noroviruses (HuNoVs)

In recent years, human noroviruses have been recognised as a cause of gastroenteritis

disease worldwide and are commonly associated with acute gastroenteritis outbreaks (Glass

et al., 2009; Patel et al., 2009). Outbreaks mainly occur in enclosed settings such as hospitals,

schools, nursing homes as well as cruise ships or military barracks (Bert et al., 2014;

Donaldson et al., 2008; Godoy et al., 2014; Hansen et al., 2007; Iturriza-Gomara & Lopman,

2014; Simon et al., 2006; Widdowson et al., 2004; Xue et al., 2014). HuNoVs are responsible

for the winter vomiting disease (also named stomach flu or gastric flu) which happens more

commonly in winter and affects people of all ages but has a more deleterious effect on

children, elderly and immune-compromised people (Estes et al., 2006; Green, 2014; Payne et

al., 2013; Rackoff et al., 2013). Each year, HuNoVs are responsible for 50% of all

gastroenteritis outbreaks (over 267,000,000 infections) (Donaldson et al., 2008) and

approximately 200,000 deaths especially in children under 5 years of age and the elderly (˃65

years) (Hall et al., 2013; Patel et al., 2008).

59

Winter vomiting disease was first recognised in 1929 by Dr Zahorsky and named as a

“Hyperemesis hemis’’ or winter vomiting disease, but the causative agent for this disease was

unknown. 43 years later, Albert Kapikian investigated stool samples from a winter vomiting

disease outbreak that occurred in an elementary school in Norwalk, Ohio, USA in 1968 and

identified a virus, naming this virus Norwalk virus (a prototype of noroviruses). It was

established as the causative agent using immune-electron microscopy (IEM) to examine stool

material from adult volunteers administered with faecal filtrate from this gastroenteritis

outbreak (Kapikian et al., 1972), later the name was changed to “small round structure virus”

(Green et al., 2000). HuNoVs are found mainly in GI and GII and few GIV norovirus

genogroups (Caddy et al., 2014; Kroneman et al., 2013; Rackoff et al., 2013; Siebenga et al.,

2009; Zheng et al., 2006). The symptoms vary depending on many factors such as age but the

most common symptoms are vomiting especially in infants, and diarrhoea as well as nausea,

abdominal pain, abdominal cramps, anorexia, myalgia and low fever (Atmar & Estes, 2006;

Patel et al., 2009). Recently, the numbers of HuNoV infection have increased and HuNoVs

cause 18% of all cases of gastroenteritis and are responsible for more than 50% of all

gastroenteritis outbreaks worldwide and between 75/90% of nonbacterial gastroenteritis

outbreaks in developing countries (Ahmed et al., 2014; Hall et al., 2014; Patel et al., 2008;

Scallan et al., 2011; Siebenga et al., 2009). In the United States, the number of HuNoV cases

is between 19 and 23 million each year, leading to 56,000-71,000 hospitalisations and 570-

800 deaths which cost the government about $777 million every year (Hall et al., 2013).

Overall the cost associated with foodborne outbreaks is $2 billion (Glass et al., 2009; Hall et

al., 2013; Hoffmann et al., 2012). In the United Kingdom, annual expenses associated with

HuNoV infection are estimated to be over £100 million (Lopman et al., 2004).

The most common genotype associated with the majority (~80%) of HuNoV

gastroenteritis including food-borne outbreaks is GII.4 whereas GI which is more stable in

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water is responsible for water-borne outbreaks (Matthews et al., 2012; Patel et al., 2008;

Rackoff et al., 2013). Worldwide, 55 to 85% of cases and outbreaks from 1990s to 2013,

which included four pandemics, were caused by seven different variants of genotypes GII.4

including: the US 1995/96 variant in 1996 (White et al., 2002), the Farmington Hills variant

in 2002 (Widdowson et al., 2004), the Hunter variant in 2004 (Bull et al., 2006), the 2006a,

2006b variants in 2007-2008 (Eden et al., 2010), and the New Orleans variant from 2009 to

2012 (Yen et al., 2011). More recently, a new variant of genotype GII.4 of HuNoVs (called

Sydney2012) was reported in New South Wales, Australia (van Beek et al., 2013). This then

spread around the world to replace the previous variant of genotype GII.4 (New Orleans

2009) as a commonly detected HuNoV strain in many countries (Allen et al., 2014; da Silva

et al., 2013; Fonager et al., 2013; Leshem et al., 2013; Shen et al., 2013; van Beek et al.,

2013; Vega et al., 2014). No significant differences in the number of outbreaks between

Sydney 2012 and New Orleans 2009 strains exist, but the new strain is associated with

distinct genetic changes in the P2 component of the capsid protein VP1 sequence compared

with that of the New Orleans 2009 strain (Allen et al., 2014; Debbink et al., 2013). These

changes to six amino acid positions, including 294, 310, 359, 368, 373 and 396, play an

important role in defining histo-blood group antigens (HBGAs) leading to the antigenic

variation responsible for increasing fitness of Sydney 2012 strains. These antigenic variations

play a critical role for the emergence of a novel strain by promoting viral escape from host

immunity (Allen et al., 2014; Debbink et al., 2012).

Typically, a new variant emerges every 2-3 years to replace the previous dominant

variant of genotype (Siebenga et al., 2007, Desai et al., 2012, Gastanaduy et al., 2013). Most

variants usually emerge within the GII.4 genotype but also among GII.2 and GI.3 (Zheng et

al., 2006, Eden et al., 2013). Four factors can influence the evolution rate of NoVs; these are

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host receptor recognition, sequence space, duration of population immunity, and replication

kinetics (Bull & White, 2011).

1.9.4 Clinical signs

The incubation period of HuNoV is usually 24-48 hrs and the symptoms vary

depending on factors such as age and the infectious dose (Kaplan et al., 1982; Karst, 2010;

Murata et al., 2007; Patel et al., 2009). The illness is commonly characterised by nausea,

vomiting (more common in children below 1 year) and watery diarrhoea (more common in

adults) as well as other clinical features including anorexia and in some cases abdominal

cramps and pain, fever, headache, chills and myalgia (Deng et al., 2007; Kaplan et al., 1982;

Karst, 2010). The duration of illness is usually 2 to 4 days, but can be longer in nosocomial

outbreaks (4-6 days) (Kaplan et al., 1982; Lee et al., 2013; Lopman et al., 2009; Murata et

al., 2007; Rockx et al., 2002; Sukhrie et al., 2011) lasting up to six weeks in infants (Patel et

al., 2009).

Although, the illness is self-limiting in the majority of cases, there is increasing

evidence that norovirus may lead to post infection disorders leading to death in the elderly

and immunocompromised patients (Harris et al., 2008; Rondy et al., 2011; Sukhrie et al.,

2010). In elderly patients mortality can be associated with dehydration (Harris et al., 2008;

van Asten et al., 2011). Studies have recognised chronic infection in immunocompromised

patients (Bok & Green, 2012; Ronchetti et al., 2014; van Asten et al., 2011). HuNoV is also

associated with other important clinical diseases including necrotizing enterocolitis (Stuart et

al., 2010; Turcios-Ruiz et al., 2008) and afebrile seizure in infants (Chan et al., 2011), as well

as pneumatosis intestinalis (Kim et al., 2011) and Crohn’s disease (Cadwell et al., 2010). In

the UK, this leads to around 80 deaths annually among elderly people (older than 64 years)

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(Harris et al., 2008). In addition, according to the WHO estimations, approximately 200,000

deaths per year occur in children below 5 years of age in developing countries (Patel et al.,

2008).

Despite a short duration of virus infection, prolonged viral shedding is reported for 3-

8 weeks following infection with high viral loads (up to 1012 NoV copies/g faeces) in

symptomatic or asymptomatic patients after recovery (Atmar et al., 2008; Rockx et al., 2002)

shedding can also continue over two years in immunocompromised patients and transplant

patients (Bok & Green, 2012). Moreover, in some cases of paediatric oncology patient the

virus shedding continues for over a year (Simon et al., 2006).

1.9.5 Transmission

Three general routes are involved in HuNoV transmission including; person-to-person

transmission directly via the faecal-oral route (consumption of food or aerosolized vomitus)

or indirectly through contaminated surfaces or fomites. The second route is food-borne

transmission via an infected food handler or food distribution system contaminated with

human waste. Finally, contaminated water is a vehicle for HuNoV outbreaks (Harris et al.,

2014; Isakbaeva et al., 2005; Yoder et al., 2008; Yoder et al., 2004). The transmission of

HuNoV is facilitated by several factors including low virus infectious dose, 8-1000 virus

particles, prolonged viral shedding, environmental stability, genetic diversity and short term

immunity which all play a critical role during HuNoV outbreaks (Debbink et al., 2013;

Duizer et al., 2004a; Hall, 2012; Hansman et al., 2006; Kirkwood & Streitberg, 2008;

Siebenga et al., 2008; Simmons et al., 2013; Teunis et al., 2008; Zheng et al., 2006). The

majority of HuNoV outbreaks occur in closed and semi-closed settings such as schools,

restaurants, cruise ships, nursing homes, and hospitals. In addition to the virus characteristics:

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low infectious dose, the short incubation period, high viral load in stool and vomitus as well

as environmental stability; the transmission of NoVs in these closed settings is dependent on

other conditions such as factors associated with patients, visitors and staff behaviour (Harris

et al., 2014; Iturriza-Gomara & Lopman, 2014; Lopman et al., 2012; Patel et al., 2009).

1.10 Model systems for the study of human norovirus biology

Since its discovery in 1972, our knowledge of HuNoVs biology, pathogenesis,

immunity, translation and replication remains incomplete due to the lack of a suitable tissue

culture system or any small animal model to propagate and study these viruses (Karst et al.,

2014a; Karst et al., 2014b; Thorne & Goodfellow, 2014). So far, several attempts have been

made to propagate HuNoV in different types of tissue or cell line as well as many animal

models using a variety of methods and techniques, but with very limited success (Cheetham

et al., 2006; Duizer et al., 2004b; Papafragkou et al., 2013; Rockx et al., 2005; Straub et al.,

2011; Straub et al., 2007; Subekti et al., 2002; Takanashi et al., 2014; Wobus et al., 2006).

This impairs the efforts to design and develop drugs or vaccines against these viruses (Karst,

2010; Rocha-Pereira et al., 2014; Rohayem et al., 2010; Thorne & Goodfellow, 2014). Early

attempts used human volunteers to understand some aspects of the NoV pathogenicity as well

as protective immunity (Dolin et al., 1971; Green et al., 1993). However, there are many

limitation for using humans as a model system including difficulty and cost, and

biocontainment requirement for this type of virus (type B pathogen) (Vashist et al., 2009).

Many studies have attempted to infect non-human hosts with HuNoV using different animals

previously reported as susceptible to enteric caliciviruses such as pigs, monkeys and calves

(Vashist et al., 2009). These attempts have largely been unsuccessful, apart from some

positive results in chimpanzees, and macaques (Bok et al., 2011; Subekti et al., 2002; Wyatt

64

et al., 1978). Limited virus replication has been reported in Rhesus macaques (Rockx et al.,

2005) and in pigtail macaque (Subekti et al., 2002) infected with HuNoV. In addition, viral

replication also has been reported to occur in chimpanzees infected with HuNoV, but this led

to different clinical symptoms from those that occur in humans (Bok et al., 2011; Wyatt et

al., 1978). GII of HuNoV replication also can occur in gnotobiotic pigs and calves, leading to

viremia (Cheetham et al., 2006; Souza et al., 2008) and the majority of infected pigs develop

mild diarrhoea (Cheetham et al., 2006). Recently, (Taube et al., 2013) reported limited

replication of human NoVs using Rag-/-γc-/- BALBc deficient mice as a model, but the

infected mice did not develop clinical symptoms and the virus was cleared within 3 days. The

molecular components required for HuNoV infection, as well as the protective immunity

against HuNoV, cannot be identified using most animal models due to the difficulty in

genetically manipulating bovine, porcine and feline hosts when compared to mouse (Wobus

et al., 2006), emphasising the need to develop a tissue culture system.

To date, no cell culture systems have successfully propagated HuNoV, in spite of

several attempts using human and non-human cell culture (Duizer et al., 2004b; Papafragkou

et al., 2013). Only three studies by Straub and colleagues (Straub et al., 2011; Straub et al.,

2007; Straub et al., 2013) have had a limited success in cultivating GI and GII of human

NoVs in a 3-dimensional system using the human embryonic intestinal epithelial cell line

INT-407 and Caco-2 cells, but other workers have been unable to reproduce these findings

(Herbst-Kralovetz et al., 2013; Papafragkou et al., 2013; Takanashi et al., 2014). In other

studies, 19 cell culture models from 11 different animals were infected with a sample known

to contain HuNoV, but none of these cells were susceptible to HuNoV (Malik et al., 2005). In

addition, Duizer et al., 2004b used a variety of human and animal cell lines including, A549,

AGS, Caco-2, CCD-18, CRFK, CR-PEC, Detroit 551, Detroit 562, FRhK-4, HCT-8, HeLa,

HEC, HEp-2, Ht-29, HuTu-80, I-407, IEC-6, IEC-18, Kato-3, L20B, MA104, MDBK,

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MDCK, RD, TMK, Vero and HEK293, as well as different laboratory methods, but all these

efforts were unsuccessful to propagate HuNoV in any of these cell lines (Duizer et al.,

2004b). Recently, a low level of HuNoV infection was detected in vitro using human B cells

in cell culture, however this required co-infection with enteric bacteria and therefore may not

be an ideal system (Jones et al., 2014). Therefore, most of the studies in recent years on

HuNoV are largely dependent on surrogate models (Vashist et al., 2009).

The animal caliciviruses are used as surrogates for the study of HuNoVs include

Sapovirus, porcine enteric calicivirus (PEC), feline calicivirus and Tulane virus (Rocha-

Pereira et al., 2014). The differences between these viruses and human noroviruses in regards

to genome organization, transmission and pathogenicity make them less than ideal for the

discovery of a HuNoV vaccine (Rocha-Pereira et al., 2014), but they provide an opportunity

to understand the pathogenicity and translation/replication mechanisms of HuNoV (Chaudhry

et al., 2006; Karst, 2010; Thorne & Goodfellow, 2014; Wobus et al., 2006).

FCV is frequently used as a model for the study of human noroviruses in regard to

environmental stability, survival, diagnostic and inactivation studies (Bae & Schwab, 2008;

Bozkurt et al., 2014; Buckow et al., 2008; Topping et al., 2009). This virus is also used as a

model to study human norovirus replication and translation due to the possibility of genetic

manipulation of the FCV genome (Chaudhry et al., 2006; Willcocks et al., 2004).

Discovery of murine norovirus in 2003 (Karst et al., 2003) and subsequent

propagation of this virus in tissue culture using cells of the hematopoietic lineage such as

macrophages (MΦ) and dendritic cells (DCs) (Wobus et al., 2004) provided an efficient cell

culture model for the study of many aspects of norovirus biology (Arias et al., 2012; Wobus

et al., 2006). At present, MNV is considered the best model for HuNoV due to its genetic

similarity, protein function similarity and ability to replicate in cell culture and in small

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animal models, providing opportunities for specific genetic manipulation in the host (Karst et

al., 2003; Wobus et al., 2004; Wobus et al., 2006; Zheng et al., 2006). Therefore several

studies have used MNV as a model for understanding many aspects of HuNoVs life cycle

(Chaudhry et al., 2006; Thorne & Goodfellow, 2014; Vashist et al., 2009; Wobus et al.,

2006). MNV replication also occurs in many murine macrophage and dendritic cell-lines

including (RAW 264.7, IC21, J774A.1, P388D1, WBC264-9C and JAWSII) (Changotra et

al., 2009; Wobus et al., 2006). This raised the possibility that HuNoV might replicate in

similar cells of human origin and share a similar tropism to cells of the hematopoietic lineage

(Leung et al., 2010). However, Norwalk virus is unable to replicate in macrophage or

dendritic cells derived from the peripheral blood of susceptible humans suggesting that

human and murine norovirus tropisms differ (Lay et al., 2010). More recently, amino acid

changes in the genome of some MNV strains including MNV-3 and MNV.CR6 have been

associated with persistence and tropism (Arias et al., 2012; Nice et al., 2013). The MNV

model has provided new insight into how HuNoV interact with their host and has generated

questions regarding the relationship between noroviruses and cells of the hematopoietic

lineage and the role of innate immunity in the control of norovirus infection (Wobus et al.,

2006).

Recently, Tulane virus (TV) was isolated from stool samples of rhesus macaques, and

due to the ability to propagate this virus in tissue culture (Monkey kidney cell line LLC-

MK2), TV may provide another valuable model for understanding HuNoVs biology (Farkas

et al., 2008).

In addition to these models, the transfection of human hepatoma Huh-7 cells with

HuNoV RNA leads to virus replication but the newly formed viruses are unable to infect

other cells. This may be due to the absence of specific factors associated with viral entry

(Guix et al., 2007). A viral replicon system for NoV was generated by Chang and co-worker

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(2006) and used to study a variety of potential antiviral molecules against HuNoV infection

(Bok et al., 2008; Chang & George, 2007; Chang et al., 2006). Recombinant virus like-

particles (rVLPs) of HuNoVs generated by expression of the major capsid protein (VP1),

with or without the minor capsid protein (VP2) (Lin et al., 2014; Vashist et al., 2009), serve

as a good model to study viral attachment (Harrington et al., 2002a) and provide opportunity

to understand the immunity against HuNoVs or develop antivirals (Harrington et al., 2002b;

Vashist et al., 2009).

1.11 Pathogenesis and immunity

Studies on NoVs pathogenicity and immunity have been hampered by the lack of a

reproducible culture system or small animal model for the propagation of human norovirus.

However, studies using FCV and MNV as models, as well as information from studies using

human volunteer have provided valuable information (Karst, 2010). Recently, animals such

as pigs and calves as well as non-human primates, infected with HuNoV, were used as

experimental animal models for understanding HuNoV biology (Bok et al., 2011; Cheetham

et al., 2006; Souza et al., 2008; Souza et al., 2007). Although viral RNA is detected in sera or

cerebrospinal fluid of infected volunteers, the upper part of intestinal tract (duodenum and

upper jejunum) remains the only target for NoVs infection (Ito et al., 2006; Takanashi et al.,

2009). Proximal intestinal biopsies have revealed specific histological changes in the

intestinal epithelium including broadening and blunting of the intestinal villi, shortening of

the microvilli, crypt-cell hyperplasia, and epithelial cell disarray and cytoplasmic

vacuolisation. In addition, infiltration of inflammatory cells (polymorphonuclear and

mononuclear cells) into the lamina propria is also observed (Troeger et al., 2009). In contrast,

no histological changes can be detected in the gastric fundus, antrum and colonic mucosa

68

(Levy et al., 1976). Gastric emptying is delayed during infection and this may be responsible

for nausea and vomiting (Meeroff et al., 1980). A transient malabsorption of fat, D-xylose

and lactose is also reported during infection and associated with the alteration of brush border

enzyme activity (alkaline phosphatase, sucrase and trehalase) in infected cells (Agus et al.,

1973). The susceptibility or resistance to infections depend primarily on the presence of

specific human HBGA receptors in the human gut (Lindesmith et al., 2008). Although more

than 80% of adults appear susceptible to HuNoVs infection, some adults display resistance to

infection due to a lack of HBGA receptors (Lindesmith et al., 2003). In addition, the

expression of HBGAs correlates with strain-specific susceptibility (Lindesmith et al., 2003).

HBGAs are a family of carbohydrates expressed in the mucosal surfaces of the respiratory,

genitourinary and digestive tracts, or linked to proteins/lipids on the red blood cell surface or

present as free oligosaccharides in biological fluids such as milk and saliva (Hutson et al.,

2004). They serve as receptors for NoVs and the binding to these receptors varies depending

on the strains (Lindesmith et al., 2008). Immunological studies reported that volunteers

display short-term immunity against NoV infections, varying from 4 to 16 weeks and

potentially as long as 2 years (Atmar et al., 2011; Johnson et al., 1990; Rocha-Pereira et al.,

2014). On average infected volunteers were repeatedly susceptible to infection with the same

strain or heterologous strains six months after previous exposure (Johnson et al., 1990;

Parrino et al., 1977). The viral dose is one of the many concerns with volunteer studies.

Although HuNoV has a very low infectious dose (less than 20 viral particles) as well as high

rate of shedding in stool (108-1010 RNA copies/gram), most studies challenged the volunteers

with doses several thousand-fold times higher than the minimum capable of causing human

illness (Teunis et al., 2008). More recently, Simmons et al., 2013 suggested that the duration

of immunity to infections lasts from about 4-8 years using six different mathematical models

69

to estimate immunity duration, depending on factors such as transmission process and natural

history of HuNoV (Simmons et al., 2013).

Both types of immunity, innate and acquired immunity have been reported to

contribute towards immunity against NoV infections (Parrino et al., 1977). The early

volunteers’ studies demonstrated no association between pre-existing serum antibodies and

the susceptibility to infection with Norwalk virus in adults (Atmar et al., 2011; Dolin et al.,

1971; Graham et al., 1994; Reeck et al., 2010). Volunteers with high levels of pre-existing

antibodies were more likely to have symptomatic illness compared to those with low levels of

antibodies (Johnson et al., 1990). In contrast, high level of antibodies may reflect short-term

immunity and recent exposure to HuNoV in children (Matsui & Greenberg, 2000). Using

MNV as a model in mice lacking functional type I interferon (IFN) signalling pathways,

studies demonstrated the requirement of type I IFN immune protection against MNV

infection (Changotra et al., 2009; Karst et al., 2003; Mumphrey et al., 2007). In addition,

(Jung et al., 2012) showed that the oral administration of type 1 IFN to gnotobiotic pigs

infected with MNV led to decrease viral shedding. IFN acts via activated STATs causing a

cascade of events leading to the induction of antiviral proteins such as RNA-dependent

protein kinase (PKR) and RNase L (Samuel, 2001). A study using a Norwalk replicon system

demonstrated that norovirus replication is inhibited by type I IFN and type II IFNγ (Chang

& George, 2007; Chang et al., 2006). The role of T-cells in this process is not completely

understood, but studies have reported that HuNoV infection or vaccination using virus-like

particles (VLPs) elicit CD4+ Th1 response leading to an increase of IFNγ and IL-2 cytokine

secretion (Lindesmith et al., 2005; Lindesmith et al., 2008). Recently, Troeger et al., 2009

observed an increase in the number of intraepithelial cytotoxic T cells in the duodenum

during the first six days of infection (Troeger et al., 2009). Moreover, this study also

correlated the apoptosis of human enterocytes with HuNoVs infections. The increase in Th1

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as well as Th2 cytokines and interferons was also reported in serum and intestinal contents of

gnotobiotic pigs and calves infected with human GII.4 strain of HuNoVs (Souza et al., 2008;

Souza et al., 2007).

Despite the complex nature of protective immunity to NoVs infection, due to the

antigenic diversity of HuNoVs, and also the lack of an efficient culture model, recent

attempts to produce a vaccine against HuNoVs using virus-like particles (VLP) have shown

promise (Herbst-Kralovetz et al., 2010; Karst, 2010; Karst et al., 2014a; Rocha-Pereira et al.,

2014; Rohayem et al., 2010; Thorne & Goodfellow, 2014; Tome-Amat et al., 2014).

1.12 Viral structure, genome organisation and function of the viral proteins

Caliciviruses are small non-enveloped spherical viruses between 28-38 nm in

diameter containing a single-stranded, positive-sense, polyadenylated RNA genome, whose

organisation varies among the calicivirus genera (Figure 1.9). While Sapovirus, Nebovirus

and Lagovirus genera have a genome consisting of two open reading frames (ORF1 and

ORF2), the genomes of Vesivirus and Norovirus are mainly organised into three ORFs (Jiang

et al., 1993; Rohayem et al., 2010). The non-structural proteins in all caliciviruses are

encoded by ORF1, but the expression of the structural proteins, the major capsid protein

(VP1) and minor capsid protein (VP2) differ among the genera. In Vesivirus and Norovirus,

VP1 and VP2 are encoded by ORF2 and ORF3, respectively; whereas in Sapovirus,

Lagovirus and Nebovirus genera, VP1 and VP2 are encoded by ORF1 and ORF2,

respectively (Rohayem et al., 2010). A common feature of all caliciviruses is the lack of a

methylated cap (m7GpppN) at the 5´ terminus of the viral RNA genome. Instead, a 10-15 kDa

protein named as VPg (Virion Protein genome-linked) is attached to the 5´end and serves as a

cap substitute for initiation of viral protein translation (Burroughs & Brown, 1978; Herbert et

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al., 1996; Karst et al., 2014a; Meyers et al., 1991). The calicivirus genome is polyadenylated

at its 3´end, terminating with a poly (A) tail of 110 nucleotides (nt) (Burroughs & Brown,

1978; Farkas et al., 2008; Jiang et al., 1993; Xi et al., 1990) (Figure 2). The mature

calicivirus virions are composed of three proteins including VP1, VP2 and VPg (Sosnovtsev

et al., 2000). While VPg is found in mature virions it only functionally serves as a non-

structural protein during replication (Green, 2013).

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Figure 1.9 Organization of representative caliciviruses genome. The caliciviruses genome is

covalently linked to VPg (NS5) at the 5’ and with a poly(A) tail and is organised into 2-4 ORFs as

indicated (A, B, C, D). ORF1 is translated as a polyprotein, cleaved by the protease NS6 at

different cleavage sites to produce the non-structural proteins that are essential for viral replication

(A, B, C, and D) and VP1 (D). VP2 is encoded by ORF3 in (A, B and C) or by ORF2 in (D).

ORF2 and ORF3 are translated from a subgenomic RNA in all caliciviruses. In addition MNV (B)

has an additional alternative fourth ORF. ORF4 overlaps with ORF2 and is also translated

primarily from the subgenomic RNA into the virulence factor 1 (VF1) protein.

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Noroviruses genome range from 7.3-7.7 kb (kilobases) in size organised into three

ORFs (Thorne & Goodfellow, 2014; Zheng et al., 2006). However, some strains of MNV

contain a fourth small ORF, ORF4, which overlaps ORF2 and is encoded by a sub-genomic

RNA (sgRNA). ORF4 encodes a 214 amino acid protein called virulence factor 1 (VF1),

involved in the regulation of the host innate immunity response and apoptosis (McFadden et

al., 2011; Thackray et al., 2007). ORF1, the largest ORF (5 kb) encodes the large poly-

protein precursor (~ 200 kDa) which is self-cleaved during translation to yield six/seven non-

structural viral proteins (NS1-7) by the viral proteinase (Pro; NS6) in the following order: the

N-terminal protein (p48; NS1-2 in MNV;37-48 kDa), the 2C-like nucleoside triphosphatase,

NTPase (NS3; ~40kDa), picornavirus 3A-like protein (p22; NS4 in MNV; 20–30 kDa), VPg

(NS5), the viral 3C-like protease 3CLpro (Pro; NS6 in MNV;19 kDa) and RNA-dependent

RNA polymerase (RdRp; NS7 in MNV; ~57 kDa) (Belliot et al., 2003; Clarke & Lambden,

2000; Hardy, 2005; Seah et al., 2003; Sosnovtsev et al., 2006).

ORF2 (1.8 kb) and ORF3 (0.6 kb) encode the structural proteins, the major capsid

protein (VP1; 57 kDa) and minor capsid protein (VP2; 22 kDa) respectively (Donaldson et

al., 2008; Glass et al., 2000a; Green et al., 2000; Jiang et al., 1992). Both of them are

encoded by a sub-genomic RNA (sgRNA; ~ 2.3 kb) (Katayama et al., 2006; Neill &

Mengeling, 1988; Wobus et al., 2004). VP2 is a small protein found in only few numbers of

copies per virion with yet unknown function (Thackray et al., 2007), but some data suggest

that VP2 may play a role in increasing the stability of VP1 by protecting it from protease

degradation (Bertolotti-Ciarlet et al., 2003), or in encapsidation of the viral genome (Karst,

2010; Vongpunsawad et al., 2013). VP2 is required for the formation of stable virus-like

particles (VLPs) via its interaction with VP1 (Bertolotti-Ciarlet et al., 2002; Di Martino &

Marsilio, 2010). VP1 protein, the main capsid protein, is composed of three domains linked

74

by a flexible hinge including the N domain, the internal part of the capsid, the S domain

forming the icosahedral shell and a third variable P domain, which contains two sub-domains

P1 and P2 forming the protrusions (Allen et al., 2008; Chen et al., 2006; Prasad et al., 1999;

Tan et al., 2008) (Figure 1.10). The P2 domain is the most hypervariable region of the capsid

which is located at the outermost capsid surface and is responsible for cellular binding

(especially to HBGAs) and immune recognition (Allen et al., 2008; Cao et al., 2007;

Chakravarty et al., 2005; Chen et al., 2006; Glass et al., 2000a; Hardy, 2005; Prasad et al.,

1999; Tan et al., 2003) (Figure 1.10B and C). Furthermore, the receptor binding specificity of

HuNoV seems to be associated with the P2 sub-domain (Prasad et al., 1999; Tan et al., 2008;

Tan et al., 2003). Therefore, this subdomain represents one of the best targets for determining

the strains that correlate with outbreaks (Lindesmith et al., 2013; Sukhrie et al., 2013; Vega

et al., 2011; Xerry et al., 2009). The 3-dimensional structure of the viral capsid determined

using X-ray crystallography showed that the virion is formed of 180 molecules of the VP1

distributed into 90 dimeric capsomeres associated with a few molecules of VP2 (usually one

or two copies) (Prasad et al., 1999; Vongpunsawad et al., 2013). The capsid protein is also

critical for viral replication through a novel interaction with the RdRp (Morillo & Timenetsky

Mdo, 2011; Subba-Reddy et al., 2012).

The function of some non-structural proteins remains unknown. Some studies have

suggested that N-terminal proteins may play a role in membrane rearrangement and

intracellular protein trafficking (Ettayebi & Hardy, 2003; Fernandez-Vega et al., 2004). NS4

and the NTPase have been shown to have functional similarity to the picornaviruses 2C and

3A protease, respectively (Hyde et al., 2009; Sosnovtsev et al., 2006). NS1-2, NS3 and NS4

proteins contribute to processes essential for MNV replication such as membrane induction or

cellular manipulation (Hyde et al., 2009). The NS6 protease shares structural similarity with

the cellular chymotrypsin-like serine proteases (Blakeney et al., 2003), and share functional

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and structural properties with picornavirus 3C proteases (Leen et al., 2012). The NoVs

proteases play an essential role in the maturation of the ORF1 polyprotein precursor into

mature non-structural proteins via cleavage of the polyprotein at different cleavage sites (QG,

EG and EA), producing six active proteins (Liu et al., 1996; Sosnovtsev et al., 2006;

Takahashi et al., 2013a; Takahashi et al., 2013b). In addition, NS6 also plays a role in the

inhibition of cellular translation via cleavage of both eukaryotic initiation factor 4G and poly

(A) binding protein (critical for initiation of translation) (Kuyumcu-Martinez et al., 2004;

Zeitler et al., 2006). In FCV, p76 acts as a protease and polymerase. The first half of p76 acts

as a trypsin-like serine protease and is related to picornavirus 3C-protease, therefore it is

designated 3C-like protease, while the second half of p76 is related to picornaviruses 3D-

polymerase and acts as an RNA-dependent-RNA polymerase (RdRp). In contrast, in MNV

the p76 is separated into two proteins including NS6 which acts as protease, and NS7 that

acts as a polymerase (Sosnovtsev et al., 2006). The crystal structure of RdRp from GII

noroviruses displayed general similarity to the structural domains of other positive strand

RNA viruses, especially with RHDV (Hardy, 2005). VPg protein (NS5) was discovered in

the late 1970s and is a small 10-15 KDa protein linked to the 5´ end of genomic and sub-

genomic RNAs in all members of Caliciviridae, and similar proteins are found in other

positive sense RNA viruses such as the Picornaviridae, Potyviridae, Luteoviridae,

Comoviridae and Secoviridae, although with different functions (Burroughs & Brown, 1978;

Clarke & Lambden, 2000; Dunham et al., 1998; Herbert et al., 1996; Kneller et al., 2006;

Kozak, 1991; Sadowy et al., 2001). For instance the VPg of picornaviruses is not involved in

translation initiation (Fitzgerald & Semler, 2009; Goodfellow, 2011), but plays a role in

replication whereas the VPg of Potyvirus (Lellis et al., 2002; Leonard et al., 2000) and

caliciviruses (Daughenbaugh et al., 2006; Herbert et al., 1997) is important for protein

synthesis. (Daughenbaugh et al., 2003) showed a direct interaction between VPg and

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eukaryotic initiation factor 3 (eIF3) of HuNoV suggesting the involvement of VPg in

translation initiation. Further evidence for the role of VPg in translation initiation came from

Goodfellow et al., 2005 who reported that FCV VPg interacts with eukaryotic initiation

factor 4E (elF4E) in infected cells and that this interaction is necessary for translation of FCV

in vitro (Goodfellow et al., 2005). This interaction between VPg and eIF4E has also been

reported in MNV-1 (Daughenbaugh et al., 2006) but the role of this interaction VPg-eIF4E

in protein synthesis is less clear (Chaudhry et al., 2006; Goodfellow, 2011). The infectivity of

caliciviruses is reduced by removal of the VPg protein from 5′end of the mRNA (Chaudhry et

al., 2006; Guix et al., 2007). In addition, VPg may play a possible role in transporting the

genome to the site of negative strand synthesis (Donaldson et al., 2008).

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Figure 1.10 The structure of norovirus major capsid protein VP1. (A) The capsid

protein is encoded by open reading frame 2 (ORF2). VP1 consists of the shell (S) and

protruding domains (P), linked by a flexible hinge, and P is divided into two subdomains,

including P1 (blue) and P2 (red). (B, C, D) P2 is located at the capsid surface and is

responsible for cellular binding and immune recognition.

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1.13 Norovirus life cycle

The exact mechanisms for HuNoV translation and replication are yet to be determined

and most knowledge originates from studies of animal surrogates (Donaldson et al., 2008;

Karst et al., 2014a; Thorne & Goodfellow, 2014; Vashist et al., 2009). Norovirus replication

occurs in the cytoplasm of the host cell, possibly enterocytes (Cheetham et al., 2006; Karst,

2010). The replication cycle of noroviruses consists of several steps (Figure 1.11) beginning

with the attachment of the VP1 P2 domain to specific cellular receptors in the surface of host

cells which in the case of HuNoVs, are the HBGAs (Abrantes et al., 2012b; Hutson et al.,

2003; Marionneau et al., 2002; Tan & Jiang, 2005). Caliciviruses such as rhesus enteric

calicivirus (ReCV), RHDV and canine noroviruses also interact with their host cells via

HBGA receptors (Caddy et al., 2014; Farkas et al., 2010; Ruvoen-Clouet et al., 2000; Tian et

al., 2007). In contrast, the interaction of other members of caliciviruses with host cells

occurs via different cellular receptors. For instance, FCV attaches to the host cell membrane

via junction adhesion molecule-1 (JAM-1) (Makino et al., 2006), whereas MNV uses a

terminal sialic acid on ganglioside GD1a of a mouse macrophage, glycolipids and

glycoproteins as a receptor depending on the MNV strains (Taube et al., 2012; Taube et al.,

2009).

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Figure 1.11 Proposed replication mechanism of Human Norovirus (HuNoV). The replication

cycle can be divided into the following stages: (1) attachment of virus via P2 to the cellular

receptor, (2) entry and (3) uncoating of the viral genome, (4, 5, 6 & 7) translation of genomic RNA

into a polyprotein precursor through VPg-dependent translation (4 & 5), followed by post-

translational processing mediated by protease (Pro) to produce six non-structural proteins (6) which

are released and accumulate in replication complex (RC) where replication occurs (7), (8, 9 & 10)

RNA replication is mediated by Pol and other non-structural proteins in RC to generate anti-

genomic RNA (negative-strand RNA) (8), which used as a template for the synthesis genomic RNA

as well as sub-genomic RNA translated as structural proteins (VP1 & VP2) (9 & 10), (11) assembly

and maturation of the structural proteins as well as packaging of the VPg-linked RNA genome

occurs, followed by release and exit of the mature virion from the cell (12).

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Many studies have reported the role of specific HBGAs in attachment of different

genotypes of NoV and recombinant virus-like particles (VLPs) to epithelial host cells,

especially Caco-2 cells (Harrington et al., 2004; Hutson et al., 2003; Marionneau et al., 2002;

White et al., 1996). The viral attachment is associated with H type 1 antigen which produces

the FUT2 enzyme (Marionneau et al., 2002). Herbst-Kralovetz et al., 2013 have reported the

lack of expression of H type 1 and 2 or Lewis b antigens on the differentiation of Int-407

cells in a 3-D cell culture system, which may explain why the human macrophage and

dendritic cells were unable to replicate HuNoVs (Papafragkou et al., 2013). Moreover, the

CEP-like morphological change of the 3-D aggregates reported by Straub et al., 2007 may be

due to the presence of lipopolysaccharides or other stool-contaminating toxins (Herbst-

Kralovetz et al., 2013). Epidemiological studies have provided strong evidence for the

requirement of NoV infection for HBGAs as receptors or co-factors (Hennessy et al., 2003;

Hutson et al., 2002; Lindesmith et al., 2003). There are two binding profile groups of NoVs,

with strains that bind A/B and/or H antigens and strains binding Lewis and/or H antigens. In

addition, another group of strains does not bind to HBGAs raising the possibility that other

receptors can be used by NoVs for infection (Donaldson et al., 2008; Murakami et al., 2013).

More recently, Murakami et al., 2013 demonstrated that the binding of NoV-like particles to

human intestinal epithelial cells line (Caco-2) occurs independently of HBGAs but depends

on the state of cell differentiation. This suggested that VLPs utilize different cell receptors or

co-receptors during binding and internalization into cells (Murakami et al., 2013).

Following attachment, the viruses enter the host cells through an unknown

mechanism. In the case of MNV, internalisation occurs by a non-clathrin, non-caveolin

mediated endocytic pathway which depends on dynamin and cholesterol (Gerondopoulos et

al., 2010; Perry & Wobus, 2010). The entry of FCV (F9 strain) occurs via clathrin mediated

endocytosis in a pH-dependent manner (Kreutz & Seal, 1995; Stuart & Brown, 2006). After

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genome uncoating, and release of the viral genome into the cytoplasm of the host cell, protein

synthesis begins with recognition of the VPg-linked RNA genome by the translation

machinery of the host cells (Rohayem et al., 2010; Thorne & Goodfellow, 2014). Caliviruses

use a novel mechanism for the initiation of protein synthesis: a VPg-dependent translation

initiation involving the interaction of eukaryotic initiation factors (eIFs) with VPg protein,

acting as a cap substitute (Chaudhry et al., 2006; Daughenbaugh et al., 2003; Goodfellow et

al., 2005; Herbert et al., 1997) (Figure 1.12). These interactions lead to recruitment of the

ribosome and host translation factors to the viral RNA and thereby initiation of translation

(Chaudhry et al., 2006; Daughenbaugh et al., 2003; Goodfellow et al., 2005).

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Figure 1.12 Schematic diagram of cap and VPg-dependent translation initiation. (A) The cap-

dependent translation mechanism is dependent of the recognition of 5’ m7Gppp(N) cap structure by the

eIF4F complex consistent of the cap binding protein eIF4E, the scaffolding protein eIF4G, the RNA helicase

eIF4A and its co-factor eIF4B. Subsequently eIF4G mediates the recruitment of the 40S ribosomal subunit

and other eIFs via an interaction with eIF3, this leads to the scanning mechanism that then locates the AUG

start codon. (B) In contrast, during VPg-dependent translation, VPg acts as a cap substitute and interacts

with eIF4E, eIF4G and eIF4A to tether ribosomes to the 5’ end of the viral RNA. During FCV infection, the

interaction of VPg with eIF4E and eIF4G is required for translation while only the interaction with eIF4G is

required during MNV infection.

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Translation of ORF1 of the viral genome produces a large single polyprotein that

subsequently undergoes auto-proteolytic processing mediated by the viral protease (3CLPro or

NS6) to release mature NS proteins and their precursors (Sosnovtsev et al., 2006). Five

3CLPro cleavage sites have been reported in noroviruses and in the case of MNV-1 NS

proteins these sites include 341E|G342, 705Q|N706, 870E|G871, 994E|A995, 1177Q|G1178 (Sosnovtsev

et al., 2006). These NS proteins along with host proteins and other viral RNA molecules

assemble in intracellular membranous structures called replication complexes (RC) where

genome replication occurs (Green et al., 2002; Hyde et al., 2009; Wobus et al., 2004). In the

case of MNV, the NS proteins p48 (NS1-2) and p22 (NS4) are required to recruit host

membranes (endoplasmic reticulum (ER), Golgi complex and endosome) and induce RC

formation, providing a platform for replication (Hyde & Mackenzie, 2010; Hyde et al., 2009;

Sharp et al., 2010) via the modulation of the host cell secretory pathway (Denison et al.,

2008). In addition, HuNoVs p48 and p22 proteins (equivalent to the MNV NS1-2 and NS4

respectively) induce disassembly of the Golgi complex and inhibit protein secretion, therefore

these proteins may have a possible role in driving cellular membrane rearrangement (Ettayebi

& Hardy, 2003; Fernandez-Vega et al., 2004; Sharp et al., 2010). Within the RC, the genomic

RNA serves as a template for anti-genomic RNA (negative sense) production via a

transcriptional process mediated by the RdRp which uridylates the VPg in the presence of

polyadenylated genomic RNA leading to the production of the negative-sense strand

(Rohayem et al., 2010). This anti-genomic RNA is then used as a template for the synthesis

of new genomic RNA, which is either translated or packaged into new virions, and sub-

genomic RNA that is translated into structural proteins (VP1 and VP2). The neo-synthesized

genomes are packaged into the capsid to assemble a mature virion which is finally released

from the cell via yet unknown mechanisms (Rohayem et al., 2010).

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The exact mechanism of sgRNA production is still unclear, but in the case of MNV

two possible mechanisms have been suggested, both dependent on RdRp. The first is via

internal initiation involving the binding of the viral RdRp to the sgRNA promoter located

downstream of the sgRNA start site in the negative strand followed by synthesis of a VPg-

linked positive strand sgRNA (Miller et al., 1985). In the second model, the RdRp binds to

the 3′ end of the positive strand genomic RNA and initiates negative-strand synthesis, and

then RdRp terminates at a specific sequence located at the 3′ end of the negative strand

sgRNA leading to production of negative strand sgRNA, which then acts as a template to

produce positive strand sgRNA (Sit et al., 1998). Other studies demonstrated the role of

RdRp of HuNoV and MNV in this process by recognising and binding to a sgRNA promoter

(Lin et al., 2015). Using this sub-genomic RNA as template, the translation of VP2 happens

through termination/reinitiation process, which depends on an upstream sequence named

termination upstream ribosomal binding site (TURBS), a 40–70 nt termination upstream

ribosomal binding site (Luttermann and Meyers, 2014).

1.14 The function of RNA-protein interaction during noroviruses infections

In addition to interactions with the host translation machinery (Figure 1.12), other

cellular proteins and conserved viral RNA structures or sequences also play an important role

in many aspects of viral life cycle (Liu et al., 2009; Lopez-Manriquez et al., 2013; Simmonds

et al., 2008; Thorne & Goodfellow, 2014; Vashist et al., 2012). Many studies have reported

direct and indirect protein-protein or RNA-protein associations between virus and their host,

some of which are required for the norovirus life cycle (Chaudhry et al., 2006; Chung et al.,

2014; Goodfellow et al., 2005; Gutierrez-Escolano et al., 2000; Karakasiliotis et al., 2010;

Lopez-Manriquez et al., 2013; Vashist et al., 2012).

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VPg forms several possible interactions with elements of the translation initiation

complex, but not all these interaction required for viral translation or replication (Chaudhry et

al., 2006; Vashist et al., 2012). These studies reported a direct or indirect association between

calicivirus VPg and specific eukaryotic initiation factors such as eIF4F components (4A, 4E

and 4G) as well as eIF3 (Chaudhry et al., 2006; Daughenbaugh et al., 2003; Goodfellow et

al., 2005). For instance, in vitro studies using reticulocyte lysate have demonstrated a direct

interaction between eIF4E (cap-binding protein) and VPg during MNV infection but this

interaction is not essential for MNV translation (Chaudhry et al., 2006; Goodfellow et al.,

2005). In contrast, the same interaction is critical in the case of FCV, as translation is

inhibited in vitro either by depletion of eIF4E or inhibition of the eIF4E-eIF4G1 interaction

(Chaudhry et al., 2006). However, the eIF4G-VPg association mediated by the C-terminal

fragment of eIF4G during MNV infection is necessary for translation and efficient viral

replication (Chung et al., 2014). Furthermore, a direct interaction between VPg and eIF3 has

been reported, but the role of this interaction is unknown (Daughenbaugh et al., 2003).

Additional indirect associations between viral VPg and other cellular proteins such as

LARP1, ABCE1, DDX9, IGFBP1, eEF1A1 and different ribosomal proteins were also

reported but the role of these interactions remain unclear (Chung et al., 2014). In vitro studies

also demonstrated that the RNA helicase eIF4A is required for MNV and FCV translation for

unwinding the secondary structure in the 5’ end of RNA, and inhibiting this initiation factor

either by using a dominant-negative form of eIF4A or a small molecule inhibitor such as

hippuristanol impairs both MNV and FCV translation (Chaudhry et al., 2006; Simmonds et

al., 2008). More recently, Royall et al., 2015 demonstrated that phosphorylation of eIF4E,

occurring via activation of the MAPK pathway is required for MNV1 replication (Royall et

al., 2015).

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Vashist et al., 2012 identified approximately 150 different proteins that interact with

the 5’ and 3’ extremities of the MNV-1 genome using rabbit reticulocyte lysate, but the role

of most of these interactions remains elusive requiring further additional studies (Vashist et

al., 2012). These interactions between the 5’ and 3’ends of the viral genome with host

proteins include Lupus autoantigen (La), polypyrimidine tract binding protein (PTB) and poly

(A) binding protein (PABP) (Vashist et al., 2012) which play an important role in MNV

replication (Gutierrez-Escolano et al., 2000; Gutierrez-Escolano et al., 2003; Kuyumcu-

Martinez et al., 2004). These studies also reported the interaction of La and polypyrimidine

tract binding protein (PTB) with both 5’ and 3’extremities of the Norwalk virus genome

(Gutierrez-Escolano et al., 2000; Gutierrez-Escolano et al., 2003). La protein, which

regulates the translation of other RNA viruses such as Hepatitis C Virus (HCV) (Kumar et

al., 2013; Ray & Das, 2011), was also reported to interact with HuNoV genome (Fitzgerald

& Semler, 2009). In the case of FCV, the interaction of PTB with the 5’ end of genomic and

sub-genomic RNA, as well as its association with RC, has been demonstrated to be essential

for viral replication (Karakasiliotis et al., 2010). In contrast, the cellular proteins La, PTB and

DDX3 have a negative impact on MNV replication in vitro (Vashist et al., 2012). Because of

the role of PTB in promoting MNV translation, but reducing translation of related viruses

such as FCV, the possibility has been raised that it plays a role in switching from translation

to replication (Karakasiliotis et al., 2010; Vashist et al., 2012).

The binding of PABP to poly (A) tails at the 3´ end of mRNAs plays a key role in

mRNA translation and stability by facilitating mRNA circularization via its binding to

eIF4G (Gorgoni & Gray, 2004). This closed loop is believed to stimulate translation (Imataka

et al., 1998; Sonenberg & Hinnebusch, 2009), and is important for promoting ribosome

cycling and stimulating 60S subunit joining (Kahvejian et al., 2005). However later in

infection, the calicivirus 3CLPro inhibits cellular translation by cleaving PABP (Kuyumcu-

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Martinez et al., 2004; Willcocks et al., 2004). Recently, Lopez-Manriquez et al., 2013

identified an important role for poly(C) binding protein (PCBP) and hnRNPA1 which bind to

5’ end of MNV-1 genome promoting genome circulation and viral replication (Lopez-

Manriquez et al., 2013). These studies also demonstrated the association of the 5’ end with

PCBP-2 and hnRNPA1 or PABP with the 3’ end. The role of these interactions remains

elusive but comparison with other positive-strand RNA viruses such as poliovirus and dengue

virus suggests a possible role in the circulation of the viral genome (Vashist et al., 2012).

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1.15 Aims and objectives of the research

It is clear from the previous section that calicivirus replication is dependent on a

complex network of interactions between the viral RNA and the host translation machinery to

mediate viral protein synthesis and to regulate translation in the host. In addition, many

studies have demonstrated that viral infections are a major stress for the host cell leading to

the regulation of mRNA translation at the level of cytoplasmic RNA granules.

Therefore the following hypothesis has been formulated: calicivirus replication

regulate the host response to infection by affecting the formation or assembly of stress

granules and P-bodies.

To address this hypothesis the following objectives have been designed:

- Using cell lines supporting FCV and MNV-1 replication to study the assembly of stress

granules and P-bodies using appropriate immunofluorescence microscopy markers.

- To dissect the effect of FCV and MNV-1 infection on the assembly of stress granules.

- To dissect the effect of FCV and MNV-1 infection on the assembly of P-bodies.

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Chapter 2

General Materials and Methods

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2.1 Maintenance of cells:

2.1.1 Crandall Rees Feline Kidney (CRFK) Cells

CRFK cells were obtained from the European Collection of Cell Cultures (ECACC)

and grown in Eagle’s Minimum Essential Medium (MEM) with Earle’s salts, supplemented

with 10% (v/v) heat inactivated foetal bovine serum (FBS), penicillin/streptomycin 1% (v/v)

and non-essential amino acids 1% (v/v) (all from GibcoBRL, Life Technologies). Cells were

maintained in a 75-cm2 tissue culture flask at 37ºC with 5% CO2 (Incubator Galaxy S; Wolf

Laboratories), and passaged as follows: the medium was removed and the cells were rinsed

with 5 ml 1:10 trypsin/versene (T/V) (all from GibcoBRL life technologies). The T/V

solution was removed and cells were detached by addition of 5 ml trypsin/versene and the

flask was incubated at 37oC with 5% CO2 for 3 min. The suspension of cells and T/V was

transferred to a Universal tube containing 5 ml of 10% FBS-Eagle’s MEM and the cells were

centrifuged at 2500 rpm (Centurion) for 3.5 min. The medium was removed and the pellet

was resuspended in 6-ml 10% FBS-Eagle’s MEM. Approximately 1-1.5 ml of cell suspension

were placed in a new 75-cm2 flask containing 14 ml 10% FBS-Eagle’s MEM. The CRFK

cells were used for all experiments were obtained between passages numbers 6 -16.

2.1.2 Mouse Leukemic monocyte-macrophage Cells (RAW 264.7)

RAW 246.7 (ECACC) cells were grown in Dulbecco’s Minimum Essential Medium

(DMEM) with Earle’s salts, supplemented with 10% (v/v) FBS, penicillin/streptomycin 1%

(v/v) (all from GibcoBRL life technologies). Cells were maintained at 37ºC with 5% CO2

(Incubator Galaxy S; Wolf Laboratories), and passaged as follows: the medium was removed

and the cells were resuspended in 8 ml 10% FBS-Eagle’s DMEM in the presence of sterile

glass beads to detach the cells from the flask. Approximately 1-1.5 ml of cell suspension

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were placed in a 75-cm2 flask containing 14 ml 10% FBS-Eagle’s DMEM. The cells were

used for all experiments were obtained between passages numbers 6 -16.

2.1.3 Feline embryonic airway (FEA) cells.

FEA cells were kindly provided from Dr Alan Radford (University of Liverpool, UK)

and grown in Eagle’s MEM with Earle’s salts, supplemented with 10% heat inactivated FBS,

1% penicillin/streptomycin and 1% non-essential amino acids. Preparation of confluent

monolayers was achieved as CRFK maintenance protocol (section 2.1.1).

2.1.4 Murine microglial cells (BV-2)

Murine microglial cells (BV2) were kindly provided from Prof Ian Goodfellow

(Cambridge University, UK). The cells were cultured in DMEM supplemented with 10%

(v/v) heat inactivated FBS, 1% penicillin/streptomycin, 1% L-glutamine and 1% sodium

bicarbonate (Life Technologies). The cells were incubated at 37ºC with carbon dioxide (5%)

in a Galaxy S incubator (Wolf Laboratories). Cells reaching a confluence of 80% were

scraped from the 75-cm2 flask substrate with sterile glass beads and centrifuged at 2500 rpm

(Centurion) for 3.5 min. The supernatant was removed and the pellet was resuspended in 8 ml

of growth medium after which 1 ml of the cell suspension was transferred into a 75-cm2

vented flask containing 14 ml of growth medium and grown at 37ºC with 5% CO2 (Incubator

Galaxy S, Wolf Laboratories).

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2.1.5 Human embryonic kidney 293 (HEK293T) cells (293 cells)

293 cells were grown in DMEM containing 1000 mg/L glucose (low glucose DMEM)

supplemented with 10% (v/v) foetal calf serum (FCS), 1% penicillin and streptomycin (Life

Technologies), 2% glutamine and 1% non-essential amino acids. The cells were incubated at

37ºC with 5% CO2 and then passaged when the confluence reached 80-90%. Preparation of

confluent monolayers was achieved as follows; the media was removed and the cells washed

with 3 ml 1% trypsin/versene (T/V) to remove remaining serum. Then, cells were detached

from the flask with 5 ml of fresh T/V. The detached cells were transferred to a 25 ml

universal tube containing 5 ml of fresh growth medium and centrifuged at 2500 rpm

(Centurion) for 3.5 min. The supernatant was removed and the cell pellet resuspended in 6 ml

of growth medium. 2 ml of the cell suspension was transferred to a 75-cm2 flask containing

16 ml fresh growth medium and incubated at 37ºC with 5% CO2 (Incubator Galaxy S, Wolf

Laboratories). Cell differentiation using poly-lysine (Sigma) treatment was applied for

culturing the cells on 13-mm coverslips as fellow; coverslips in 24-wells plate were treated

with poly-lysine solution, rocked and left at RT for 5 min. Poly-lysine were then removed

and coverslips were rinsed with sterile MQ water for several time before leaving the

coverslips to dry in a sterile hood for 2h and cells were then grown on coverslips.

2.1.6 Murine macrophage J774 cells

J774 cells (from the ATCC) were maintained in DMEM medium with 10% FBS at

37°C with 5% CO2. For sub-culturing, cells with confluence of 80-90% were scraped from

the 75-cm2 flask substrate with a cells scraper (VWR), spin down at 2500 rpm (Centurion) for

3.5 min and resuspended in 6-ml of fresh growth medium. 1ml of cell suspension was added

to 75-cm2 flask containing 14 ml of fresh growth medium.

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2.1.7 THP-1 cells

THP-1 cells (from the ATCC) were grown in RPMI 1640 medium (from ATTC)

supplemented with 2 mM L-glutamine, 0.05 mM beta-mercaptoethanol and 10% FBS. When

the cells were approximately 80% confluent, they were harvested, centrifuged at 1500 rpm

(Centurion) for 5 min and then suspended in 5-ml of fresh growth medium. I-ml of cell

suspension was added to 19-ml of growth medium and incubated at 37ºC and 5% CO2. Cell

were differentiated using PMA treatment prior immunofluorescence studies as fellow; Stock

of 5 mg/ml of PMA was prepared in DMSO and 4 µl of this stock was added to T-75 flask

containing 2-4 X 105 cells in 20 ml of media (5mg/ml). Flasks were then incubated overnight

in 37C and 5% CO2. For 3 days, following incubation, differentiated cells were washed with

PBS (3X). The PBS were then replaced by growth media.

2.2 Freezing and Resuscitation of CRFK and RAW264.7 cells

Freezing medium, 10 ml of appropriate growth medium for CRFK or RAW264.7

cells, 1 ml FBS and 1 ml dimethylsulphoxide (DMSO) was prepared for freezing cells as

follows: cells were harvested from one or more 75-cm2 flasks and subjected to centrifugation

at 2500 rpm (Centurion) for 3-4 min. The supernatant was removed and the pellet

resuspended in 2 ml freezing medium per flask and stored as 1ml aliquots in cryovials. The

cryovials were then placed overnight at -800C to freeze slowly. Next day, the cells were

transferred to liquid nitrogen storage.

For resuscitation, cells were thawed either at room temperature or rapidly in a 37ºC

water bath. The cells were then transferred to a universal tube containing 5 ml of the

appropriate warm (37ºC) medium and subjected to centrifugation at 2500 rpm (Centurion) for

3-4 min. The supernatant was discarded and the pellet resuspended in 7 ml of the appropriate

94

medium and transferred to a small flask (25-cm2). Following overnight incubation at 37ºC

with 5% CO2 (Incubator Galaxy S; Wolf Laboratories), the medium was exchanged for new

growth medium to remove any residual DMSO and the cells were incubated until confluent.

They were then passaged as normal (see section 2.1.1 and 2.1.2).

2.3 Preparation of viruses stock

2.3.1 Preparation of Feline Calicivirus (FCV) stock

A 175-cm2 flask of cells (80% confluent) was infected with 400µl of FCV Urbana

(obtained from Prof Ian Goodfellow, Cambridge University, UK) in 4 ml 2% (v/v) FBS-

Eagle’s MEM and rocked for 1 hour at room temperature (RT). After rocking, the medium

was removed and replaced with 8 ml of 10% (v/v) FBS-Eagle’s MEM and the cells incubated

at 37ºC for a further 5 hours before leaving the cells to freeze overnight at -80oC. The cells

were then thawed and scraped off and placed in a 20 ml universal, pelleted at 2500 rpm

(Centurion) for 2.5 min and the supernatant filtered through a 0.2 µm Millex filter, divided

into aliquots and stored at -80ºC. The virus titre was estimated by determination of the 50%

tissue culture infectious dose (TCID50) (Section 2.4).

2.3.2 Production of Murine Norovirus (MNV) stock

A confluent (approximately 80%) 75-cm2 flask of RAW264.7 cells was infected with

1-ml of MNV-1 (obtained from Prof H. Virgin, Washington University, St Louis, USA) seed

stock in 4 ml 2% (v/v) FBS-Eagle’s DMEM and rocked for 1h at RT. Another 8 ml of 10%

FCS- Eagle’s DMEM were added to the flask and incubated for 17 h until the cytopathic

effect was evident. Cells were frozen at -80oC overnight before being allowed to thaw at

room temperature (RT) for 1 h. The cells were subjected to a second freeze-thaw (3-5h) at -

80ºC and then centrifuged at 2500 rpm (Centurion) for 2.5 min to remove the cell debris. The

95

supernatant was filtered through a 0.2 µm Millex filter, divided into aliquots and stored at -

80oC. The viral titre was estimated by determination of TCID50 (Section 2.4).

2.4 50% Tissue Culture Infectious Dose (TCID50) Assay

TCID50 values for both FCV and MNV-1 were determined as follows: nine serial

dilutions (10-1 – 10-9) of stock virus were prepared in serum-free medium (MEM for FCV and

DMEM for MNV), 50 µl of each dilution were added to six wells of a 96-wells round-bottom

plate. One row was used as a control (no virus). Cells were prepared at a concentration of 4 x

105 cells/ml. One-hundred microliters of cell suspension were added to each well and the

cells were incubated at 37ºC with 5% CO2 overnight for FCV or 48 h for MNV. After

incubation, each well was scored as infected (positive) or uninfected (negative) and the titre

of the virus was calculated using the following formula from the method of Reed and

Muench.

Viral titre = [(% of wells infected at dilution above 50%) - 50%] .

[(% of wells infected above 50%) – (% of wells infected below 50%)]

2.5 Preparation of VPg-linked FCV RNA

Four 175-cm2 flasks containing of CRFK cells (80-100% confluent) in 30 ml medium

were used for infection with FCV. 20 ml of medium was removed from the cells and

collected in a sterile bottle. The cells were infected in the remaining 10 ml with FCV Urbana

(MOI=3) and left to rock for 30 min at RT. 5 ml of CRFK medium was added to the flask and

the cells were incubated for 6 hours at 37ºC with 5% CO2. The infected cells were scraped

off using cell scraper and decanted into 20 ml Universals. The cells were pelleted at low spin

speed (700 rpm) (Centurion) for 5 min. The supernatant was removed and filtered through a

0.2 µm Millex filter, aliquoted and stored at -80ºC. The cell pellets were pooled and washed

96

in 10 ml of PBS and spin at 2500 rpm (500g; Centurion) for 5 min. The cells were

resuspended in 4 ml of cold TN buffer (10 mM Tris [pH 7.8], 10 mM NaCl) and allowed to

swell on ice for 15 min. The cells were lysed with 60 strokes in a cold 2 ml tight-fitting glass

Dounce homogenizer (Bellco Glass, Inc. Vineland, N.J.). The lysate was transferred to four

1.5-ml Eppendorf tubes and centrifuged for 5 min at 3450 rpm (800g) at 4ºC (Micro 22R,

Hettich) to remove nuclei and un-lysed cells. The supernatants were then transferred to a

fresh Eppendorf tube and subjected to centrifugation for 20 min at 15000 rpm at 4ºC (Micro

22R, Hettich). The resulting pellets corresponding to replication complexes were each

resuspended in 120 µl of TN buffer containing 15% glycerol (replication complexes) and

stored at -80ºC. The replication complexes were used to make RNA at a later date.

2.6 Preparation of VPg-linked MNV RNA

RAW264.7 cells from (approximately 80–100 %) were infected with 1 ml of MNV-1

stock in 10 ml of FBS-Eagle’s DMEM and rocked for 1h. A further twenty ml of FBS-

Eagle’s DMEM were added to the flask and incubated for 23h at 37ºC with 5% CO2 until the

cytopathic effect was evident. After incubation the cells were scraped off and pelleted at 2500

rpm (Centurion) for 3 min. The supernatant was removed and filtered through a 0.2 µm

Millex filter, aliquotted and stored at -80ºC. The cell pellets were resuspended in 1 ml lysis

buffer supplemented with β-mercaptoethanol (GenElute kit Sigma).

2.7 Preparation and quantitation of VPg-linked FCV and MNV RNA

The GenElute Mammalian Total RNA Miniprep Kit (Sigma) was used to prepare

VPg-linked RNA from replication complexes as follows: 500µl of lysis buffer (2-ml lysis

buffer and 20µl 2-mercaptoethanol; prepared in the fume hood) was containing 250µl of

replication complexes mixed gently. The 750µl solution were then transferred into 4 blue

97

filter tubes and subjected to centrifugation at full speed for 1 min. The supernatant was

retained and equal volume of 70% ethanol was added prior to mixing. Lysate/ethanol was

applied to RNA-binding column in two aliquots and washed with 500µl of wash 1. The filter

of RNA-binding column was transferred into new tube and wash twice with 500µl of wash 2.

The columns were transferred into new tubes and 50µl of elution buffer was added. The tubes

were incubated at room temperature for a few minutes then spun at full speed (Boeco) for 1

min and the elution was collected and stored at -80ºC. One microliter of elution was used to

determine the concentration of the RNA using a Nano Drop ND-1000 spectrophotometer.

2.8 Immunofluorescence studies

2.8.1 Cell culture and virus infection

Cell of all types were maintained as previously described in Section 2.1. Cells were

seeded on 13-mm glass coverslips (VWR) at a required density overnight and virus infection

were carried out at a multiplicity of infection (MOI) of 0.2 for FCV or 30 for MNV at

different time points: 0, 2, 4, 6 and 8h and 0, 4, 8, 12, 16, 18 h, for FCV and MNV-1

respectively, unless otherwise specified. In all immunofluorescence studies, virus growth

medium was identical to the maintenance medium, except for the use of FBS at a

concentration of 2%. Mock-treated cells received 2% FBS growth medium only.

Optimization of MOI and concentration of PBS used were determined empirically during this

study.

2.8.2 Immunofluorescence assay

For immunofluorescence, cells were grown on 13-mm glass coverslips and following

chemical, physical treatment and infection, cells were fixed in 800 µl 4% paraformaldehyde

98

(Sigma Aldrich) in PBS (Appendix 1) on ice for 1h, then maintained in PBS at 4ºC until

staining. For staining, fixed cells were permeabilised by incubating with 0.1% Triton X-100

in PBS (BDH LTD) (Appendix 1) for 15 min, and then washed once with PBS. Fixed cells

were then blocked in PBS/BSA, PBS containing 1% bovine serum albumin (BSA) (Sigma

Aldrich) (Appendix 1) for 30 min. Primary and secondary antibodies (Table 2.1) were

diluted in blocking buffer. Following blocking, primary antibodies (200 µl) were added to

each well and incubated for 1 h, washed three times with PBS (3 x 5 minute washes), before

incubating for an additional 1 h with 200 µl secondary antibodies. A further 3X washes were

given before ToPro3 nuclear stain (1:10000 in PBS; Molecular Probes, Invitrogen) was

applied for 10 min. The coverslips were removed, rinsed in MQ water and placed cell-side

down on a glass slide with Vectashield mounting media (Vector Laboratories) then sealed

with clear nail varnish. All staining steps were performed at room temperature (RT). A Zeiss

LSM 510 META confocal laser microscope was used to detect SGs or PBs during infection

or chemical/physical treatment. The qualification of SGs and PBs formed were carried out

using the Image J software to determine the number of SG or PB-positive cells.

2.9 Effect of calicivirus infection on SGs and PBs formation

The MOI of the viruses used during this study was determined empirically using

confocal microscopy by calculating the percentage of infected cells as compared to the total

number of cells at different MOI. In addition, the dilutions of viral antibodies, SGs markers

proteins and PBs markers proteins used during this study were also selected empirically using

Confocal microscope (Table 2.1).

99

Figure 2.1 Primary and secondary antibodies that are used in

immunofluorescence studies

Primary antibody Company

and reference

number

Dilution Secondary antibodies Company

and reference

number

Dilution

Stress granules markers

G3BP1 BD

611126

1:600 Alexa-Anti-mouse 555 Invitrogen

A21424

1:400

PABP-1 Santa Cruz

Sc-32318

1:200 Alexa-Anti-mouse 555 Invitrogen

A21424

1:400

TIA-1 Santa Cruz

Sc-34739

and

Proteintech

17649-1-AP

12133-2-AP

Donkey anti-goat 555 Invitrogen

A21432

and

Santa Cruz

Sc-2024

1:400

eIF4G Santa Cruz

Sc-9601

Anti-mouse 555 Invitrogen

A21424

1:400

eIF3η Santa Cruz

Sc-16377

Donkey anti-goat 555 Invitrogen

A21432

1:400

P-bodies Markers

Dcp1 Santa Cruz

Sc-22574

1:50 Alexa-Anti-mouse 555 Invitrogen

A21424

1:400

Xrn-I Santa Cruz

Sc-165984

Alexa-Anti-mouse 555 Invitrogen

A21424

1:400

Viral markers

1G9 1:10000 Alexa-Anti-rabbit 488 Invitrogen

A11034

1:400

P76 1:600 Alexa-Anti-rabbit 488 Invitrogen

A11034

1:400

3D-polymerase

(NS7)

1:1500 Alexa-Anti-rabbit 488 Invitrogen

A11034

1:400

100

2.9.1 Staining for FCV-p76 protein

To visualise FCV infection, CRFK cells were stained with different dilution of anti

FCV p76 primary antibody (1:600; optimising during this study); Alexa-Fluor®-labelled anti-

rabbit secondary antibody (1:400; Molecular Probes, Invitrogen) and ToPro3 staining of

nuclei.

2.9.2 Staining for MNV-3D polymerase protein

To visualise MNV-1 infection, RAW264.7 cells were stained with different dilution

of anti MNV-3D-polymerase primary antibody (1:600; optimising during this study); Alexa-

Fluor®-labelled anti-rabbit secondary antibody (1:400; Molecular Probes, Invitrogen) and

ToPro3 staining of nuclei.

2.9.3 Staining for SG-markers

To visualise SGs formation during calicivirus infection, CRFK or RAW264.7 cells

were stained with different dilution of anti G3BP1, PABP-1, eIF4G, eIF3η, and TIA-1

primary antibodies; Alexa-Fluor®-labelled anti-mouse secondary antibody (1:400; Molecular

Probes, Invitrogen) for G3BP1, PABP, and eIF4G and donkey anti-goat IgG secondary

antibody (1:400; Molecular Probes, Invitrogen) for TIA-1 and eIF3ŋ. ToPro3 was used for

staining of nuclei.

101

2.9.4 Staining for PBs-markers

To visualise PBs formation during calicivirus infection, CRFK or RAW264.7 cells

were stained with different concentration of anti Dcp1 and Xrn1 primary antibodies; Alexa-

Fluor®-labelled anti-mouse secondary antibody (1:400; Molecular Probes, Invitrogen) and

ToPro3 staining of nuclei.

2.10 SGs formation during FCV infection

CRFK cells as well as FEA cells were plated on 13-mm coverslips slides (VWR) in

24 well plates at a concentration of 4 x 104 per well and maintained at 37°C with 5% CO2

overnight as described previously (Section 2.1.1 & 2.1.2). The next day, cells were infected

with 2% FBS MEM containing FCV at MOI of 0.2 at different time points (0, 2, 4, 6, and 8

h), the plates were rocked for 5 min and incubated at 37°C with 5% CO2 for 1 h. Mock-

treated cells received 2% FBS MEM only. Following incubation, viruses were removed and

replaced by 200 µl of 2% FBS MEM medium and the plates were incubated at 37°C with 5%

CO2 for additional 5 h. Cells were fixed for 1 h in 4% paraformaldehyde on ice for 1h, then

maintained in PBS at 4ºC until staining. The infected and mock infected cells were stained

for SGs markers (G3BP1, PABP-1, TIA-1, eIF3η and eIF4G) and p76 as a marker for virus

as described previously (Section 2.8.2).

2.10.1 Treatment with sodium arsenite (NaAsO2)

To select the optimal time of NaAsO2 (Sigma) (Appendix 1) treatment, CRFK cells

were grown on coverslips overnight prior treated the cells with 0.5mM NaAsO2 for different

times (30 min, 1h and 2h), then cell were fixed with 4% paraformaldehyde prior

102

immunostained for G3BP1 for detection the SGs using immunofluorescence microscopy as

described in Section 2.8.2.

2.10.2 Effect of FCV infection on SGs formation

CRFK and FEA cells were grown on 13-mm coverslips in 24-wells plate overnight

and then infected with FCV as above described for 6 h. Cells were then treated with 0.5 mM

NaAsO2 (Sigma) for 1h and then immunostained for G3BP1, PABP-1, TIA-1, eIF3η , eIF4G

and p76. Uninfected cells were treated with 0.5 mM NaAsO2 for 1h, infected cells as well as

untreated uninfected cells were used for comparison. The effect of FCV infection on SGs

formation was analysed using immunofluorescence microscopy as described in Section 2.8.2.

2.10.3 Effect of NaAsO2–induced SGs on FCV replication

CRFK cells were grown on 35-mm dishes overnight then treated with 0.5 mM

NaAsO2 for 1h, prior infected with FCV at different time points (0, 2, 4, 6, 8, 12, 16, and

18h). Cells supernatant then collected and used to determine the viral titre by TCID50

(Section 2.4) compared with supernatant of untreated infected cells. In addition, cell

supernatants at different time points were incubated with 13-mm coverslips containing CRFK

cells, fixed and immunostained for G3BP1 and p76 as described in Section 2.8.2.

2.10.4 Effect of FCV infection on hydrogen peroxide (H2O2) induced SGs

CRFK cells were grown on 13-mm coverslips and infected with FCV as above

described for 6 h. Cells were then treated with 1 mM H2O2 (Sigma) (Appendix 1) for 2h.

Uninfected cells were treated with 1 mM H2O2 for 2h, untreated cells and cell only infected

103

with FCV were used for comparison. Cells were stained for G3BP1 and p76 as a marker for

SGs and virus respectively. Immunofluorescence microscopy was used to determine the

effect of FCV infection on SGs formation as described in Section 2.8.2.

2.10.5 Effect of the H2O2-induced SGs on FCV replication

CRFK cells were plated on 35-mm dishes overnight, treated with 1 mM H2O2 for 2h,

and then infected with FCV at different time points (Section 2.8.1). Cells supernatant then

collected at each time point and used to determine the viral titre using TCID50 (Section 2.4).

In addition, cells at different time points were immunostained for G3BP1 and p76 to visualise

the SGs and virus as described in Section 2.8.2.

2.11 Preparation of FCV free supernatants

2.11.1 Virus Infection and clarification (virus elimination)

CRFK cells were seeded in 35-mm dishes at concentration of 2 x 105 cells and

incubated overnight. On the following day, the cells were infected with FCV for 6h, the

supernatant were then collected, transferred to Eppendorf tubes and spun at 4000 rpm at 4ºC

(Micro 22R, Hettich) for 30 min. After centrifugation, supernatant was filtered through a 0.2

µm Millex filter (Figure 2.2).

104

Figure 2.2 Preparation of the FCV-free supernatant from FCV-infected cells for

immunofluorescence studies.

105

2.11.2 Virus Precipitation

To precipitate the virus, solid NaCl was added to the supernatant at a final

concentration of 0.2M whilst mixing in cold room. Then, PEG3350 was slowly added to a

final concentration of 10%. The sample was then incubated onto a rotating wheel and mixed

overnight at 4ºC. The following morning, the samples were centrifuged for 60 minutes at

14000 rpm (Micro 22R, Hettich) at 4ºC. The supernatants were transferred to ultracentrifuge

tubes and span at 50000 rpm for 2h at 4ºC using SW55Ti rotor (Beckman). The supernatant

was then filtered through a 0.2 µm Millex filter, divided into aliquots (100µl each), and

stored in -20ºC until required.

2.13 Virus inactivation using ultraviolet-light (UV-light)

FCV pools or supernatant were UV inactivated at a wavelength of 254 nm for 4 min

(3 times) using a cross linker (Stratalinker® UV Crosslinker) before being used to inoculate a

wells of a 24-well plate containing CRFK cells. The cell were infected with FCV or FCV

supernatant and incubated at 37 ºC with 5% CO2 in a Galaxy S Incubator for 1 and 6 h. Cells

were then fixed using 4% paraformaldehyde for 1h prior to maintenance with PBS in 4ºC

until immunofluorescence study were carried out. In addition, the virus titre (with or without

UV treatment) were determined using TCID50 (Section 2.4) to confirm UV inactivation of the

virus.

106

2.14 Effect of cell culture supernatant on stress granules formation

CRFK, FEA, RAW264.7, BV2, 293T, J744, and THP-1 cells were grown on 13-mm

coverslips overnight before incubated for 6 h with the supernatant of FCV-infected or mock-

infected. Cells were then fixed and stained for G3BP1 and p76. Untreated CRFK cells were

incubated with solution of NaCl and PEG3350 at the same concentration as used in virus

precipitation protocol and followed by staining for G3BP1 to detect any effect on SGs

induction as described in Section 2.8.2.

2.15 Effect of interferon on stress granules formation

CRFK cells were grown on 13-mm coverslips overnight before incubating with

different concentration (150, 200, 300, and 3000 IU/ml) of interferon α (IFN α) for 6 h. Stress

granules were identified by immunofluorescence microscopy as described in Section 2.8.2.

2.16 Effect of RNAse A treatment on stress granules formation

RNAse A was added to FCV-infected CRFK supernatant (1U/1000 µl), incubated at

37oC for 15 min. CRFK cells were cultured on coverslips in 24-well plate overnight and then

treated with 200 µl of supernatant containing RNAse A. Control cells were incubated with

untreated supernatant. Cells were rocked for 10-15 min, incubated at 37ºC with 5% CO2 for 6

h, fixed using 4% paraformaldehyde for 1h and then immunostained for G3BP1 and p76 for

immunofluorescence microscopy as described in Section 2.8.2.

107

2.16 Effect of heat shock on stress granules formation

FCV-infected cells supernatant and mock-infected supernatant were incubated at 65ºC

for 10 and 20 min. 200µl of pre-heated supernatant was incubated with CRFK cells grown on

13-mm coverslips in 24-wells as previous described, rocked for 10-20 min and incubate at

37ºC with 5% CO2 for 6 h. Cells were then fixed with 4% paraformaldehyde for 1h and

stained for G3BP1 and p76 for immunofluorescence microscopy as described in Section

2.8.2.

2.17 SGs formation during MNV infection

RAW264.7 cells as well as BV-2 cells were cultured overnight as previously

mentioned (Section 2.1.2 &2.1.3). The following day, cells were plated on 13-mm coverslips

slides in 24-well plates at a concentration of 4.5 x 104 per well and incubated at 37°C with

5% CO2 overnight. The following day, cells were infected with MNV-1 at MOI of 30 for

different time (0, 4, 8, 12, 16 and 18 h). The plates were rocked for minutes and incubated at

37°C with 5% CO2 for 1 h. Mock-treated cells received media only. Following incubation,

the media were removed and replaced by 200 µl of 2% FBS MEM medium and the plates

were incubated at 37°C with 5% CO2. Cells were then fixed using 4% paraformaldehyde on

ice for 1h, then maintained in PBS at 4°C until staining. MNV-1 and mock infected cells

were stained for SGs markers (G3BP1, TIA-1, and eIF3η) and 3D-polymerase as a marker

for virus for immunofluorescence microscopy as described in Section 2.8.2.

108

2.18 Formation of P-bodies during FCV Infection.

CRFK and FEA Cells were seeded in 13-mm coverslips (VWR) in 24-well plates at a

concentration of 4 x 104 per well and maintained at 37°C with 5% CO2 overnight as

previously described. Next morning, cells were then infected with FCV at MOI 0.2 at

different time (0, 2, 4, 6 and 8h), cells then fixed after each time points with 4%

paraformaldehyde for 1h and maintained in PBS until staining for PBs markers. Cells were

then stained for Dcp1 and Xrn1 as PB markers as well as for p76 as virus marker for

immunofluorescence microscopy as described in Section 2.8.2.

2.19 Formation of P-bodies during MNV Infection.

RAW264.7 and BV2 cells were seeded in 13-mm coverslips (VWR) in 24-well plates

at a concentration of 4.5 x 104 per well and maintained at 37°C with 5% CO2 overnight as

previously described. Next morning, cells were infected with MNV-1 at MOI 30 for different

time (0, 4, 8, 12, and 18h), cells were then fixed after each time points with 4%

paraformaldehyde for 1h and maintained in PBS until staining for PB markers. Cells were

then stained for Dcp1and Xrn1 as PB markers as well as for 3D-polymerase as virus markers

for immunofluorescence microscopy as described in Section 2.8.2.

2.20 Statistical analysis

Experiments were carried out in triplicate with two samples each time, except where

specified. GraphPad Prism 6 was used for statistical testing using one-way ANOVA and

Dunnets post-doc testing to compare more than 2 samples or t-tests for comparing two

samples. Statistical significance was considered as P < 0.05. Image J software was used for

109

quantification of cells displayed stress granules or P-bodies during calicivirus infection. More

than 10 fields were randomly chosen and counted, with at least 400 cells counted for each

sample slide.

110

Chapter 3

Regulation of Stress Granule

formation during FCV and MNV-1

infection

111

3.1 Optimisation of experimental conditions for infection and SGs detection

To control the host response to infection and allow viral persistence, many viruses have

the ability to modulate the formation of cytoplasmic RNA granules in infected cells (Reineke

& Lloyd, 2013; Tsai & Lloyd, 2014). To date, the role played by RNA granules in the virus-

host relationship mediated by caliciviruses remains unknown. Therefore, to gain insights into

how caliciviruses modulate host stress responses during infection, I used FCV and MNV-1 as

surrogate models for human norovirus, and examined their effect on SG formation. As this was

the first project to investigate this line of research in our laboratory, a large body of my initial

work focussed on the optimisation of experimental conditions for immunofluorescence

detection of viruses and choice of marker proteins for the detection of SG assembly.

First, to determine the optimal MOI required for getting appropriate infection, I infected

the CRFK cells for 6h with different FCV MOI, from 0.1 to 1, and RAW264.7 cells for 12h

with different MNV MOI from 0.2 to 30. Cells were then fixed and stained for FCV VP1, the

capsid protein, or MNV NS7, the pol protein (see introduction, section 1.12), prior to indirect

immunofluorescence assay using confocal microscopy to determine the proportion of infected

cells (Figure 3.1 & 3.2). Among the different MOI tested, the best results were obtained at an

MOI of 0.2 for FCV and 30 for MNV where between 65-75% of the cells were found to be

infected (Figure 3.1C & 3.2K). These MOI were adopted for all subsequent experiments as

they allowed investigation of the formation of cytoplasmic granules in both infected and

neighbouring bystander cells during calicivirus infection.

Several dilutions of antibodies directed against p76, the pol-pro fusion protein, from

1:300 to 1:600 were then used to identify the optimal antibody concentration yielding strong

and specific labelling with low background noise in CRFK cells. As seen in Figure 3.3, the p76

112

dilution 1:600 gave an optimal signal: noise ratio for the detection of viral particles and this

dilution was then used in all subsequent studies.

113

ToPro-3

ToPro-3

ToPro-3

P76

P76

IG9

P76

Overlay

Overlay

Overlay

Figure 3.1 Optimization of FCV MOI for immunofluorescence studies. CRFK cells

were infected for 6h at several MOI as indicated (B-F), then cells were fixed and stained

for FCV infection using 1G9 antibody (green; middle panels) followed by Alexa-488

secondary antibody. ToPro3 stained nuclei are shown in blue (left panels). Right panels

show an overlay of both antibodies. Cells were visualised using a ZEISS LSM META

confocal laser microscope. The bar indicates a size of 10 µm.

A

B

C

D

E

F

- FCV

FCV

MO1 0.1

FCV

MOI 0.2

FCV

MOI 0.3

FCV

MOI 0.6

FCV

MOI 1

1G9

ToPro-3 1G9 Overlay

ToPro-3 1G9 Overlay

1G9

ToPro-3 Overlay 1G9

114

-MNV-1

MNV-1

MOI 0.2

MNV-1

MOI 0.5

MNV-1

MOI 1

MNV-1

MOI 2

MNV-1

MOI 5

MNV-1

MOI 10

A

B

C

D

E

F

G

ToPro-3 Overlay NS7

ToPro-3 Overlay NS7

Overlay NS7 ToPro-3

Overlay NS7 ToPro-3

Overlay NS7 ToPro-3

Overlay NS7 ToPro-3

Overlay NS7 ToPro-3

- Continued -

115

Figure 3.2 Optimization of MNV-1 MOI for immunofluorescence studies.

RAW cells were infected for 12h at several MOI as indicated (B-K), then cells

were fixed and stained for MNV-1 infection using NS7 antibody (green, middle

panels) followed by Alexa-488 secondary antibody. ToPro3 stained nuclei are

shown in blue (left panels) and the right panels show an overlay of both antibodies.

Cells were visualised using a ZEISS LSM META confocal laser microscope. The

bar indicates a size of 10 µm.

MNV-1

MOI 15

MNV-1

MOI 20

MNV-1

MOI 25

MNV-1

MOI 30

H

I

J

K

ToPro-3 Overlay NS7

ToPro-3 Overlay NS7

ToPro-3 Overlay NS7

ToPro-3 Overlay NS7

116

Figure 3.3 Optimization of p76 antibody dilution for immunofluorescence

studies. CRFK cells were infected for 6h at MOI of 0.2, then infected (B-E) and

uninfected (A) cells were fixed and stained for FCV infection using several dilution

of p76 antibody (B-D) followed by Alexa-488 secondary antibody (green). ToPro-3

stained nuclei are shown in blue (left panels) and middle panels show p76 while the

right panels show an overlay of both antibodies. Cells were visualised using a ZEISS

LSM META confocal laser microscope. The bar indicates a size of 10 µm.

ToPro-3

ToPro-3

ToPro-3

ToPro-3

p76

p76

p76

p76

Overlay

Overlay

Overlay

Overlay

A

B

C

D

E

- FCV

FCV

1:300

FCV

1:400

FCV

1:500

FCV

1:600

Overlay p76 ToPro-3

117

3.2 Chemical induction of SGs using oxidative stress in CRFK cells

SG formation is induced by a wide range of experimental stresses such as heat shock

(Anderson & Kedersha, 2002; Kedersha et al., 1999; Nover et al., 1983), UV irradiation

(Anderson & Kedersha, 2006; Moutaoufik et al., 2014), hypoxia(Anderson & Kedersha, 2006;

Takahashi et al., 2013), exposure to sodium arsenite (Anderson & Kedersha, 2006; Khaperskyy

et al., 2014), hydrogen peroxide (Emara et al., 2012) and Pateamine A (Anderson & Kedersha,

2009b; Dang et al., 2006; Khaperskyy et al., 2014). These are useful tools to dissect whether

stress granules are assembled in response to stresses communicated in an eIF2-dependent or

independent manner.

The most commonly used SG inducer is sodium arsenite treatment which results in the

phosphorylation of eIF2α via the HRI kinase (Anderson & Kedersha, 2009b; Kedersha &

Anderson, 2007; Kedersha et al., 1999; McEwen et al., 2005). To assay the ability of CRFK

cells, supporting FCV replication, to respond to stress and form SGs, we treated these cells

with 0.5 mM of sodium arsenite for 30 min, 1h or 2h and subsequently monitored the assembly

of SGs using G3BP1 as marker (Figure 3.4). The immunofluorescence analysis revealed that

1h was the optimal length of treatment for induction of SGs, reflected in the strong

accumulation of cytoplasmic G3BP1 foci, while 30 min did not result in any foci formation,

and 2h led to abnormal morphology and death of most of the cells (Figure 3.4D).

118

Figure 3.4 Induction of stress granules by oxidative stress. CRFK cells were

incubated with 0.5 mM sodium arsenite (SA) for various time points as mediated

on the panels B-D, then cells were fixed and stained for G3BP1 antibody (red;

middle panels), followed by Alexa-555 secondary antibody. ToPro-3 stained

nuclei are shown in blue. Cells were visualised using a ZEISS LSM META

confocal laser microscope. The bar indicates a size of 10 µm.

A

B

C

D

- SA

+ SA

30 min

+ SA

1h

+ SA

2h

ToPro-3 G3BP1 Overlay

ToPro-3 G3BP1 Overlay

ToPro-3 G3BP1 Overlay

ToPro-3 G3BP-1 Overlay

119

3.3 Optimisation of the markers and conditions used for stress granule detection in CRFK

cells

To monitor SG assembly during infection and not rely on only one marker, G3BP1, we

assayed the potential use of diverse SG markers in CRFK cells. Based on informal discussion

with Dr Dominique Weill in Université Paris 6 (Paris, France), we opted for the following

potential stress granule markers: G3BP1, TIA-1, PABP, eIF4G, eIF3. CRFK cells were

stressed with sodium arsenite (0.5 mM 1h) and the assembly of SGs was monitored by

immunofluorescence microscopy using G3BP1, PABP, TIA-1, eIF3η and eIF4G, using

unstressed cells as control. While sodium arsenite treatment resulted in the accumulation of

G3BP1 and PABP-1 cytoplasmic foci (Figure 3.5 A & B), no signal could be detected for

eIF4G, eIF3η and TIA-1 (Figure 3.5C, D & E). For all antibodies, a large range of dilutions

was used to identify the most suitable dilution for SG detection (data not shown).

120

eIF4G eIF4G

eIF3η

TIA-1

TIA-1

eIF3η

G3BP1 G3BP1

PABP-1 PABP-1

eIF3η

TIA-1

G3BP1

Figure 3.5 Detection of stress granules by oxidative stress in FCV-infected cells

using various markers. CRFK cells were incubated with 0.5 mM sodium arsenite (SA)

for 1h, treated and untreated cells were then fixed and stained for (A) G3BP1, (B)

PABP-1, (C) eIF4G or (D) eIF3η and (E) TIA-1 followed by Alexa-555 secondary

antibody. ToPro3 stained nuclei are shown in blue. Cells were visualised using a ZEISS

LSM META confocal laser microscope. The bar indicates a size of 10 µm.

- SA + SA

A

B

C

D

E

G3BP1

121

The amount of serum used in the cell culture media is a key component of a successful

infection but can also act as a source of stress on cells. For instance, serum-starvation is a potent

inducer of SG formation in certain cell types used for cell culture systems (Anderson &

Kedersha, 2009b; Dewey et al., 2011; Kedersha et al., 1999). The use of serum and glucose-

free medium can induce SG assembly in HepG2 hepatocytes cells (Hanson & Mair, 2014).

The common protocol for FCV infection recommends the use of serum-free media,

which could potentially induce stress, therefore I analyzed the effect of fetal bovine serum

(FBS) concentration on SG assembly by incubating CRFK cells with different concentrations

of FBS (0, 2, 5 and 10%) and monitored the impact on SGs by immunofluorescence microscopy

using G3BP1 as SG marker (Figure 3.6). While the absence of serum induced the formation of

stress granules, the lowest FBS concentration tested, 2%, did not and was selected for the

subsequent infections.

122

ToPro-3 G3BP1 Overlay

ToPro-3 G3BP1 Overlay

ToPro-3 G3BP1 Overlay

ToPro-3 G3BP1

0% FBS

2% FBS

5% FBS

10% FBS

A

B

C

D

Overlay

Figure 3.6 Optimization of FBS concentration for detect of stress granules in CRFK

cells. CRFK cells were grown overnight using different concentration of FBS as indicated

(A-D), cells were then fixed and stained for G3BP1 antibody (red; middle panels), followed

by Alexa-555 secondary antibody. ToPro3 stained nuclei are shown in blue. Cells were

visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size

of 10 µm.

123

3.4 Optimisation of the markers used for stress granule detection in RAW264.7 cells

To monitor SG assembly in a cell line supporting MNV-1 infection, we stressed

RAW264.7 cells with sodium arsenite (0.5 mM 1h) and used immunofluorescence microscopy

to detect G3BP1, TIA-1 and eIF3, using unstressed cells as control. Unexpectedly, none of

the above markers displayed aggregation into cytoplasmic foci following sodium arsenite

treatment (Figure 3.7). Several different antibody dilutions were tested without any further

success (not shown). These results suggest that RAW264.7 may be intrinsically unable to

assemble SGs, at least in response to oxidative stress.

124

G3BP1

TIA-1

eIF3η eIF3η

TIA-1

G3BP1

Figure 3.7 Detection of stress granules in RAW264.7 cells using various markers.

RAW264.7 cells were treated with 0.5mM SA , then fixed and stained for (A) G3BP1 or

(B) TIA-1 and (C) eIF3η followed by Alexa-555 secondary antibody. ToPro3 stained nuclei

are shown in blue. Cells were visualised using a ZEISS LSM META confocal laser

microscope. The bar indicates a size of 10 µm.

- SA + SA

A

B

C

125

3.5 Stress granule formation during FCV infection.

To investigate the effect of FCV infection on the formation of SGs and whether

cytoplasmic granules are part of the response to the virus, CRFK cells were infected with FCV

Urbana at MOI 0.2 for 8h, and the presence of SGs was investigated by immunofluorescence

assay microscopy at several time points: 0h, 2h, 4h, 6h and 8h using G3BP1 as marker for

stress granules and p76 as marker for viral particles (Figure 3.8). For each time point in the

FCV-infected or mock-infected cells, the number of cells displaying SGs were quantified and

the experiments performed in triplicate (Figure 3.9).

The experiments revealed that in mock-infected cells only very few cells were able to

form SGs over the 8h time course. Overall, SGs were only detected at 4, 6 and 8h post-infection

(Figure 3.14 panels C, D and E). The induction was modest was a maximum of 15% of cells

displaying SGs at 4h post-infection, against 1.4% for mock-infected cells. A closer inspection

of the results actually indicates that although overall the infection results in an increase in the

number of cells displaying SGs, we could not detect any SGs in cells infected with FCV (see

Figure 3.8D for a close up view at 6h post-infection). All SGs formed are detected in

uninfected, neighbouring or bystander cells that are in the vicinity of infected cells (Figure 3.8).

In summary these experiments suggest that FCV infection does not induce SG formation in

infected cells, while SG formation was induced in uninfected cells located near foci of

infection.

126

Figure 3.8 Detection of stress granules during FCV infection. CRFK cells were infected

with FCV at MOI of 0.2 for several time points as mediated on the panels B-E, and then

immunostained for G3BP1 antibody (red) and p76 (green) as markers for stress granules

and virus respectively followed by Alexa-555 and Alexa-488 secondary antibodies. Left

panels show p76 alone (green), middle panels show G3BP1 alone (red) and right panels

show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). On panel D a close-

up view is displayed on the side. Cells were visualised using a ZEISS LSM META confocal

laser microscope. The bar indicates a size of 10 µm.

p76 G3BP1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

A

B

C

D

E

- FCV

FCV 2hpi

2HPI

FCV 4hpi

2HPI

FCV 6hpi

2HPI

FCV 8hpi

2HPI

Overlay G3BP1 p76

127

h o u rs p o s t- in fe c t io n

Ce

lls

dis

pla

yin

g

str

es

s g

ra

nu

les

(%

)

0 2 4 6 8

0

5

1 0

1 5

2 0

2 5

- F C V

+ F C V

ns

ns

**

****

Figure 3.9 Quantification of the number of cells displaying stress granules during FCV

infection. CRFK were infected with FCV at MOI of 0.2 for several time points, stained for

G3BP1 and p76 as described in Figure 3.8. Confocal images were then used to quantify the

numbers of cells with stress granules using Image J software. Results shown are the

percentage of cells displayed SGs at each time point compared to the control, and are the

mean of three independent experiments (+/-SE; ** is P ≤0.01; ns not significant).

128

To confirm these results, the 6h time point was repeated using PABP-1 as a marker for

SGs. As shown in Figure 3.10, the infection also resulted in the induction of stress granules,

with 8% of cells displaying SGs during infection as compared to 3.9% for mock infected, but

this difference was not found to be significant after statistical analysis (Figure 3.11). However,

SGs were only detected in neighbouring cells close to infection foci and none could be detected

within FCV-infected cells (Figure 3.10).

Furthermore, to confirm that this effect was not cell-line specific, we infected feline

embryonic fibroblast (FEA) cells, that support FCV replication, with FCV for 6h and monitored

SG assembly using G3BP1 as marker for SGs and p76 as marker for the virus. Similarly to

results obtained with CRFK cells, infection of FEA cells with FCV did not result in SG

induction, and only uninfected cells displayed SGs (Figure 3.12).

129

PABP-1

PABP-1 p76

p76 Overlay

Overlay

Figure 3.10 Detection of stress granules during FCV infection using PABP-1 as a

marker. CRFK cells were infected with FCV at a MOI of 0.2 for 6h, then infected and mock

cells immunostained for PABP-1 (red) and p76 (green) as markers for stress granules and

virus respectively followed by Alexa-555 and Alexa-488 secondary antibodies. From the

left, the second panels show p76 alone (green), third panels show PABP-1 alone (red) and

right panels show an overlay of both. Left panels show cell nuclei were stained with ToPro-

3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The

bar indicates a size of 10 µm.

- FCV

+ FCV

6hpi

130

- F

C V

+ F

CV

6h

0

5

1 0

1 5

2 0

2 5C

ell

s d

isp

lay

ing

str

es

s g

ra

nu

les

(%

)

n s

Figure 3.11 Quantification of the number of cells displaying stress granules during

FCV infection using PABP-1. CRFK were infected with FCV at MOI of 0.2 for 6h, stained

for PABP-1 and p76 as described in Figure 3.10. Confocal images were then used to quantify

the numbers of cells with stress granules using Image J software. Results show are the

percentage of cells displayed SGs at compared to the control, and are the mean of three

independent experiments (+/-SE; ns: not significant).

131

p76 G3BP1 Overlay

p76 G3BP1 Overlay

Figure 3.12 Detection of SGs in FEA cells during FCV infection. FEA cells were infected

with FCV at MOI of 0.2 for 6h, and then immunostained for G3BP1 antibody (red) and p76

(green) as markers for SG and virus respectively. Left panels show p76 alone (green), middle

panels show G3BP1 alone (red) and right panels show an overlay of both. Cell nuclei were

stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser

microscope. The bar indicates a size of 10 µm.

- FCV

+ FCV

6hpi

132

The previous optimisation section revealed that none of the following markers for SGs,

TIA-1, eIF4G or eIF3 allowed successful detection of SGs in unstressed cells. However, we

cannot exclude that this marker could accumulate in the granules detected during infection, in

neighbouring cells. To test for this, CRFK cells infected with FCV for 4h, which displayed SGs

as revealed using G3BP1, were assayed by immunofluorescence microscopy using TIA-1 or

eIF3 as stress granule markers. As shown in Figure 3.13, no cytoplasmic foci could be

detected with these markers, confirming that they do not allow successful detection of SGs in

CRFK cells, infected or stressed with sodium arsenite.

133

ToPro-3 p76 Overlay TIA-1

ToPro

-3 p76 Overlay TIA-1

p76 eIF3η overlay ToPro-3

p76 eIF3η overlay ToPro-3

p76 eIF4G overlay ToPro-3

A

B

+ FCV

p76 eIF4G overlay ToPro-3

- FCV

- FCV

+ FCV

- FCV

+ FCV

Figure 3.13 Detection of SGs formation during FCV infection using different markers.

CRFK cells were infected for 6h at MOI of 0.2, then uninfected and infected cells were

fixed and stained for FCV infection using p76 antibody (green) and for different SG marker

including (A) TIA-1, (B) eIF3η and (C) eIF4G antibody (red) followed by Alexa-488 and

Alexa-555 secondary antibodies. ToPro3 stained nuclei are shown in blue. Cells were

visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size

of 10 µm.

C

134

3.6 Stress granule formation during MNV-1 infection.

To investigate whether the absence of SGs was a pattern that was conserved in other

caliciviruses, I examined the formation of SGs during MNV-1 infection. While RAW264.7

cells do not form SGs in response to sodium arsenite treatment, we could not exclude that they

would respond to stress mediated by MNV-1 infection. To test this, RAW264.7 cells were

infected with MNV-1 at an MOI of 30 for 18h, and examined by immunofluorescence

microscopy at 0, 4, 8, 12 and 18h post-infection using G3BP-1 as a marker for SGs, and NS7

as marker for MNV-1 (Figure 3.14). MNV-1 infection did not result in the assembly of

cytoplasmic G3BP-1 foci at any of these times (Figure 3.14).

To confirm that this complete absence of SGs, unlike during FCV infection, was not

due to a cell-specific or marker-specific effect, RAW264.7 cells were also examined using

TIA-1 and eIF3η as SG markers at 12h post-infection (Figure 3.15). MNV-1 infection did not

result in assembly of cytoplasmic foci for any of these markers (Figure 3.15). Next, BV2

murine microglial cells, also supporting MNV-1 infection, were infected with MNV-1 for 12h

or stressed with sodium arsenite (0.5 mM 1h) and examined using G3BP1, TIA-1 and eIF3η

as SG markers (Figure 3.16). None of these experimental conditions led to the detection of SGs

during infection, and only G3BP1 accumulated in SGs following sodium arsenite stress.

Overall, this lead to the conclusion that cells that support MNV-1 infection do not

respond to sodium arsenite treatment or infection by assembling canonical stress granules and

therefore subsequent studies focussed only on the effect of FCV.

135

- SA/- MNV-1

+ SA/- MNV-1

+ MNV-1

0hpi

+ MNV-1

4hpi

+ MNV-1

8hpi

+ MNV-1

12hpi

+ MNV-1

18hpi

G3BP1 NS7 Overlay

G3BP1 NS7 Overlay

G3BP1 NS7 Overlay

G3BP1 NS7 Overlay

G3BP1 NS7 Overlay

G3BP1 NS7 Overlay

G3BP1 Overlay

Figure 3.14 Detection of stress granules during MNV-1 infection. RAW264.7 cells were infected

with MNV-1 at a MOI of 30 for different time points or treated with sodium arsenite (SA) as indicated

(B-G) and then immunostained for G3BP1 (red) and NS7 (green) as markers for stress granules and

virus respectively, followed by Alexa-555 and Alexa-488 secondary antibodies. Left panels show

NS7 alone (green), middle panels show G3BP alone (red) and right panels show an overlay of both.

Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META

confocal laser microscope. The bar indicates a size of 10 µm.

NS7

A

B

C

D

E

F

G

136

- MNV-1

- MNV-1

+ MNV-1

+ MNV-1

TIA-1

eIF3η

Figure 3.15 Detection of stress granules during MNV-1 infection using various markers.

RAW264.7 cells were infected with MNV-1 for 12h, infected and mock cells were then fixed and

stained for (A) TIA-1 and (B) eIF3η followed by donkey anti goat secondary antibody. ToPro3 stained

nuclei are shown in blue. Cells were visualised using a ZEISS LSM META confocal laser microscope.

The bar indicates a size of 10 µm.

Overlay TIA-1 p76

Overlay TIA-1 p76

Overlay eIF3η p76

Overlay eIF3η p76

A

B

137

eIF3η

TIA-1

- SA/- MNV-1

+ SA/- MNV-1

+ MNV-1

12hpi

A

B

eIF3η Overlay p76

eIF3η Overlay p76

eIF3η Overlay p76

TIA-1 Overlay p76

TIA-1 Overlay p76

p76 TIA-1 Overlay

- SA/- MNV-1

+ SA/- MNV-1

+ MNV-1

12hpi

138

G3BP1

Figure 3.16 Detection of stress granules during MNV-1 infection in BV-2 cells using different

markers. BV-2 cells were infected with MNV-1 at a MOI of 30 for 12h or treated with sodium

arsenite (SA; 1h 0.5mM) and then immunostained for various markers for SG (A-C), and for MNV

infection using NS7 antibody. Left panels show NS7 alone (green), middle panels show G3BP1

alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue).

Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a

size of 10 µm.

C

Overlay G3BP1 p76

Overlay G3BP1 p76

Overlay G3BP1 p76

- SA/- MNV-1

+ SA/- MNV-1

+ MNV-1

12hpi

139

3.7 FCV infection impairs the assembly of stress granules induced by sodium arsenite.

It is well documented that the detection of viral dsRNA, formed as intermediate during

viral RNA replication, is a major stress for the host cell and a potent activator of stress pathways

resulting in SG formation (Dewey et al., 2011; Okonski & Samuel, 2013; Reineke & Lloyd,

2013; Takahashi et al., 2013; Zhang et al., 2014). The previous results in Figure 3.8, suggested

that FCV infection might actively prevent the formation of SGs during infection.

Therefore to dissect this further, we analysed whether FCV-infection could prevent the

formation of SGs induced by sodium arsenite. To this end, CRFK cells were either mock or

FCV-infected for 2h or 6h, and then were treated with 0.5 mM sodium arsenite for 1h to induce

the formation of SGs. The ability of CRFK cells to form SGs was then addressed by

immunofluorescence using G3BP1 as the SG marker. As shown on Figure 3.17, mock-infected

cells that are treated with sodium arsenite are competent in assembling SGs with 83.4% of cells

displaying G3BP1 foci (Figure 3.17, Figure 3.18). However, when cells were FCV-infected

prior to treatment with sodium arsenite the ability to assemble SGs was impaired with only 7%

of cells containing G3BP1 foci at 6h post-infection (Figure 3.17, Figure 3.18). In contrast, at

2h post-infection there is less effect of FCV infection on the ability of CRFK cells to assemble

SGs with 41% of cells displaying SGs (Figure 3.18). These results strongly suggest that FCV-

infection impairs the ability of cells to assemble stress granules in response to sodium arsenite,

and that this effect is more pronounced later in the replication cycle.

140

overlay G3BP1 p76

overlay G3BP1 p76

Figure 3.17 Detection of stress granules induced by NaAsO2 following FCV infection. CRFK cells were infected with FCV for 2h and 6h, treated with sodium arsenite (SA) to induce stress granule formation after each time for 1h and then immunostained for G3BP1 (red) and p76 (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle panels show G3BP alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.

-FCV/-SA

-FCV/+SA

+FCV 6hpi/-SA

+FCV 6hpi/+SA

overlay p76 G3BP1

overlay G3BP1 p76

overlay

overlay G3BP1 p76

G3BP1 p76

+FCV 2hpi/-SA

+FCV 2hpi/+SA

overlay

overlay

G3BP1 p76

p76

141

Ce

lls

dis

pla

yin

g

str

es

s g

ran

ule

s (

%)

-FC

V/-

SA

-FC

V/+

SA

+F

CV

2h

/-S

A

+F

CV

2h

/+S

A

+F

CV

6h

/-S

A

+F

CV

6h

/+S

A

0

2 0

4 0

6 0

8 0

1 0 0*

**

Figure 3.18 Quantification of the number of cells displaying stress granules induced

by NaAsO2 following FCV infection at different time points. CRFK were infected with

FCV at MOI of 0.2 for 2h and 6h, stained for G3BP1 and p76 as described in Figure 3.17.

Confocal images were then used to quantify the numbers of cells with stress granules using

Image J software. Results shown are the percentage of cells displaying SGs at each time

point compared to the control, and are the mean of three independent experiments (+/-SE;

* is P ≤0.05, ** is P≤0.01 for 2h and 6h respectively).

142

We further confirmed these results by using another SG marker, PABP, and the same

experimental design. As shown on Figure 3.19, mock-infected cells respond to sodium arsenite

treatment with 71.7% of cells assembling PABP cytoplasmic foci (Figure 3.20). In contrast,

only 21% of FCV-infected cells that are treated with sodium arsenite could assemble SGs

(Figure 3.19, Figure 3.20). Although the inhibition of SG induction was not as strong as when

using G3BP1 as marker, this confirms that FCV infection impairs the ability of cells to

assemble SGs in response to oxidative stress induced by sodium arsenite.

143

overlay PABP-1 p76

overlay PABP-1 p76

A

B

C

D

overlay PABP-1 p76

overlay PABP-1 p76

-SA/-FCV

+SA/-FCV

-SA/+FCV 6hpi

+SA/+FCV 6hpi

Figure 3.19 Detection of stress granules induced by sodium arsenite (SA) following

FCV infection using PABP-1 as a marker. CRFK cells were infected with FCV for 6h,

treated with sodium arsenite (SA; 0.5mM 1h) to induce stress granule formation and then

immunostained for PABP-1 (red) and p76 (green) as markers for stress granule and virus

respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone

(green), middle panels show PABP-1 alone (red) and right panels show an overlay of both.

Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM

META confocal laser microscope. The bar indicates a size of 10 µm.

144

- F

CV

-S

A

- F

CV

+S

A

+ F

CV

-S

A

+ F

CV

+S

A

0

2 0

4 0

6 0

8 0

1 0 0

Ce

lls

dis

pla

yin

g

PA

BP

-1 c

on

tain

ing

-

SG

s (

%)

****

Figure 3.20 Quantification of the number of cells displaying stress granules induced by

sodium arsenite following FCV infection using PABP-1 as a marker. CRFK were

infected with FCV at MOI of 0.2 for 6hpi, stained PABP-1 and p76 as described in Figure

3.19. Confocal images were then used to quantify the numbers of cells with stress granules

using image J software. Results shown are the percentage of cells displayed SGs compared

to the control, and are the mean of three independent experiments (+/-SE; **** is P

≤0.0001).

145

3.8 FCV infection impairs the assembly of stress granules induced by hydrogen peroxide.

Several stress pathways are responsible for the induction of SGs in cells (Anderson &

Kedersha, 2009a; b; Mazroui et al., 2006; Montero & Trujillo-Alonso, 2011; Tsai & Lloyd,

2014). The results from section 3.7 suggest that FCV infection prevents the induction of SGs

that are mediated in an eIF2 phosphorylation dependent manner. SG assembly can however

also be induced by an eIF2 phosphorylation independent pathway (Dang et al., 2006; Emara

et al., 2012; Mazroui et al., 2006). For example, the induction of SGs by Pateamine A and

hippuristanol is not dependent on the phosphorylation of eIF2 but instead occurs through an

effect on the function of the RNA helicase eIF4A, impairing eIF4F activity and translation

(Dang et al., 2006; Mazroui et al., 2006). Furthermore, the induction of SGs by hydrogen

peroxide is mediated by the inhibition of the cap-binding eIF4F complex rather than through

the phosphorylation of eIF2α. Hydrogen peroxide treatment promotes the hypo-

phosphorylation of 4E-BP1 resulting in the sequestration of eIF4E and disruption of eIF4F

assembly (Emara et al., 2012).

Therefore to assess whether FCV infection could prevent the induction of SGs in an

eIF2 phosphorylation independent manner, we used hydrogen peroxide as a SG inducer.

CRFK cells were either mock or FCV-infected for 6h, and then were treated with 1 mM

hydrogen peroxide for 2h to induce the formation of SGs. The ability of CRFK cells to form

SGs was then analysed by immunofluorescence using G3BP1 as the SG marker. As shown in

Figure 3.21, mock-infected cells that are treated with hydrogen peroxide are competent in

assembling SGs with 87.7% of cells displaying G3BP1 foci (Figure 3.21, Figure 3.22).

However, when cells were FCV-infected prior to treatment with hydrogen peroxide, the ability

to assemble SGs was impaired with only 7.7% of cells containing G3BP1 foci (Figure 3.21,

146

Figure 3.22). These results strongly suggest that FCV infection impairs the ability of cells to

assemble stress granules in response to hydrogen peroxide and therefore in an eIF2

phosphorylation independent manner.

147

-FCV/-H2O2

-FCV/+H2O2

+FCV/-H2O2

+FCV/+H2O2

overlay G3BP1 p76

overlay G3BP1 p76

overlay G3BP1 p76

overlay G3BP1 p76

Figure 3.21 Detection of stress granules induced by hydrogen peroxide (H2O2) following FCV infection. CRFK cells were infected with FCV for 6h, treated with hydrogen (H2O2) to induce stress granule formation and then immunostained for G3BP1 antibody (red) and p76 antibody (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle panels show G3BP1 alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.

148

- F

CV

- H

2O

2

+ H

20

2

+ F

CV

+ F

CV

+ H

20

2

0

2 0

4 0

6 0

8 0

1 0 0

Ce

lls

dis

pla

yin

g

str

es

s g

ran

ule

s (

%)

***

Figure 3.22 Quantification of the number of cells displaying stress granules induced by

hydrogen peroxide (H2O2). CRFK were infected with FCV at MOI of 0.2 for 6hpi, stained

for G3BP1 and p76 as described in Figure 3.21. Confocal images were then used to quantify

the numbers of cells with stress granules using image J software. Results shown are the

percentage of cells displayed SGs compared to the control, and are the mean of three

independent experiments (+/-SE; *** is P ≤0.001).

149

3.9 Effect of Stress granule induction on FCV replication

Overall, FCV infection prevents the assembly of SGs induced by stress signalling via

both the eIF2-dependent and independent pathways. This raises the possibility that FCV

replication impairs the assembly of SGs to prevent the formation of intracellular compartments

that would putatively have an antiviral effect on the propagation of the virus.

To test this hypothesis, we monitored whether chemical induction of stress granules,

prior to infection with FCV, would alter viral replication. To this end, CRFK cells were treated

with either sodium arsenite or hydrogen peroxide to induce SG formation and the replication

of FCV was determined by measuring viral titres at 6 or 8h post-infection using a TCID50 assay.

As shown in Figure 3.23, the induction of SGs, either with sodium arsenite or hydrogen

peroxide impaired viral replication with reduction in titre from 1,61.10+6 to 0 TCID50/ml for

both treatment at 6h post-infection, and 6,21.10+6 to 0 TCID50/ml at 8h post-infection, for both

treatments. Therefore FCV infection may impair SG formation to prevent an antiviral response

from being triggered.

150

6h u

ntreate

d

6h S

A

6h H

2O

2

8h u

ntreate

d

8h S

A

8h H

2O

2

1 0 -1

1 0 0

1 0 1

1 0 2

1 0 3

1 0 4

1 0 5

1 0 6

1 0 7

TC

ID5

0/m

L

F C V in fe c t io n (h p i)

Figure 3.23 Effect of stress granules induction on FCV replication. CRFK cells were

treated with 0.5 mM sodium arsenite (SA) for 1h or 1 mM hydrogen peroxide (H2O2) for

2h, then infected with FCV at MOI of 0.2 for 6h (black) or 8h (grey). The supernatants of

infected cells and treated infected cells for each time-point were used to determination of

viral titre by TCID50. Results show viral titre from treated cells in each time point compared

to the viral titre from untreated infected cells, and are the mean of three independent

experiments.

151

3.10 Effect of FCV infection on pre-assembled stress granules

The results from sections 3.4, 3.7 and 3.8 strongly suggest that FCV infection is actively

impairing the formation of SGs as a response to various stresses. However, we also asked the

question whether FCV infection could induce the dissolution of pre-existing SGs. Upon

treatment with sodium arsenite, cells have the ability to assemble SGs. Following removal of

the stress, cells then recover while SGs are dissolved and many factors are suggested to be

involved in this process such as turnover of SG protein component, chaperone proteins, and

autophagy (Buchan, 2014). In addition, tyrosine-phosphorylation-regulated kinase 3 (DYRK3)

plays a possible role in stress granule dissolution through control of mTORC1 signalling during

cellular stress (Wippich et al., 2013). Furthermore, the disassembly of SGs upon recovery may

be due to dephosphorylation of eIF2α (Kedersha et al., 1999).

Therefore to test the ability of FCV infection to dissolve pre-existing SGs, CRFK cells

were treated with sodium arsenite for one hour then mock- or FCV-infected and the number of

cells displaying stress granules monitored over an 8h time course (Figure 3.24). Following the

removal of sodium arsenite, mock-infected cells quickly recovered from the stress, with the

number of cells displaying SGs dropping from 83% to 65% by 2h post-infection, then 32% by

4h post-infection and continuously dropping further. At similar times during FCV infection,

the number of cells displaying SGs were 20.8% and 43.2% at 0 and 2h post-infection

respectively and no stress granules could be detected at later time points (Figure 3.25).

Therefore this could suggest that pre-assembled SGs disappeared more quickly during FCV

infection when compared to a mock infection, and could reflect the fact that FCV infection

could induce the disassembly of pre-existing stress granules. However these results are difficult

to interpret as one could also argue that two mechanisms are overlapping: the inhibition of

152

stress assembly through an inhibition of stress signalling by FCV infection, and the active

dissolution of stress granules.

153

-Continued-

0hpi

2hpi

-SA/-FCV

+SA/-FCV

0.5 mM NaSAO2

+ FCV

p76 G3BP1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

+SA/+FCV

0.5 mM NaSAO2

+ FCV +SA/-FCV

0.5 mM NaSAO2

+ FCV +SA/+FCV

0.5 mM NaSAO2

+ FCV

154

6hpi

+SA/+FCV

0.5 mM NaSAO2

+ FCV

4hpi

Figure 3.24 Effect of FCV infection on pre-assembled stress granules. CRFK cells were

treated with 0.5 mM sodium arsenite (SA) for 1h, then infected with FCV at MOI of 0.2 for

several time points as noted on the panels. Cells were then immunostained for G3BP1

antibody (red) and p76 (green) as markers for stress granules and virus respectively,

followed by Alexa-555 and Alexa-488 secondary antibodies. Left panels show p76 alone

(green), middle panels show G3BP1 alone (red) and right panels show an overlay of both.

Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM

META confocal laser microscope. The bar indicates a size of 10 µm.

p76 G3BP-1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

p76 G3BP1 Overlay

+SA/-FCV

0.5 mM NaSAO2

+ FCV +SA/+FCV

0.5 mM NaSAO2

+ FCV +SA/-FCV

0.5 mM NaSAO2

+ FCV

155

F C V in fe c tio n (h p i)

Ce

lls

dis

pla

yin

g

str

es

s g

ran

ule

s (

%)

0 2 4 6 8

0

2 0

4 0

6 0

8 0

1 0 0

-S A /-F C V

-S A /+ F C V

+ S A /-F C V

+ S A /+ F C V

Figure 3.25 Quantification of the number of cells displaying stress granules upon sodium

arsenite (SA) treatment and FCV infection. CRFK were treated with 0.5 mM SA for 1h prior

infected with FCV at MOI of 0.2 for several time-points, stained for G3BP1 and p76 as

described in Figure 3.26. Confocal images were then used to quantify the numbers of cells with

stress granules using Image J software. Results shown are the percentage of cells displayed

SGs compared to the untreated uninfected, SA-treated and untreated infected, and are

representative of one replicate.

156

3.11 Effect of UV-inactivation on the inhibition of stress granule assembly

Because infection of CRFK cells with FCV led to the inhibition of SG formation

induced in both an eIF2 phosphorylation dependent and independent manner, this raised the

question whether this inhibition could be due to active viral replication or a cascade of cellular

events that would follow viral attachment.

To test these possibilities we used UV inactivation of FCV virus particles. Viral

particles were exposed to UV light (3 pulses of 4 min at 120,000 microjoules). To confirm the

inactivation, the ability of untreated and UV-inactivated viruses to replicate was then

determined using TCID50 assay. As shown in Figure 3.26, UV-inactivation strongly impaired

the ability of FCV to replicate with a reduction in titre from 1,28.10+6 to 0 TCID50/ml.

Subsequently untreated and UV-inactivated FCV particles were used to infect CRFK cells, and

6h post-infection these cells were challenged with sodium arsenite to induce SG assembly. The

formation of stress granules was monitored by immunofluorescence using G3BP1 as a marker

(Figure 3.27). As seen in Figure 3.27, UV-inactivation of FCV did not impair the induction of

SGs following sodium arsenite treatment, with 85.8% of cells displaying SGs as compared to

15.5% for untreated FCV particles (Figure 3.28). Therefore this strongly suggests that in order

to impair the assembly of stress granules, FCV particles need to enter the infected cells and

undergo active replication.

157

FC

V 6

h

UV

-in

act i

vate

d F

CV

6h

1 0 -1

1 0 0

1 0 1

1 0 2

1 0 3

1 0 4

1 0 5

1 0 6

1 0 7

TC

ID5

0/m

L

**

Figure 3.26 Effect of UV-inactivation on FCV replication. FCV were exposed to UV-light

as described in materials and methods, then incubated with CRFK cells for 6h. The viral titre

was determination using TCID50. Results shown are the titre of UV-inactivated virus compared

to the viral titre of active FCV, and are the mean of three independent experiments (+/-SE; **

is P ≤0.01).

158

-FCV/-SA

-FCV/+SA

+FCV 6hpi/

-SA

+FCV (UV) 6hpi/

-SA

+FCV (UV) 6hpi/

+SA

Figure 3.27 Effect of UV-inactivation on the inhibition of stress granules assembly. CRFK cells were infected with FCV or UV-inactivated FCV for 6h, treated with sodium arsenite (SA) to induce stress granule formation. Infected cells (C, D & E) and Mock cells (A & B) were then immunostained for G3BP1 (red) and p76 (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle panels show G3BP-1 alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.

A

B

C

D

E

overlay G3BP1 p76

overlay G3BP1 p76

overlay G3BP1 p76

overlay G3BP1 p76

overlay G3BP1 p76

159

-FC

V/-

SA

-FC

V/+

SA

+F

CV

(U

V)/

-SA

+F

CV

(U

V)/

+S

A

+F

CV

/+S

A

0

2 0

4 0

6 0

8 0

1 0 0

Ce

lls

dis

pla

yin

g

str

es

s g

ra

nu

les

(%

)

**

n s

Figure 3.28 Quantification of the number of cells displaying stress granules during

FCV or UV-inactivated FCV infection and sodium arsenite (SA) treatment. CRFK were

infected with FCV at MOI of 0.2 for 6h, stained for G3BP1 and p76 as described in Figure

3.22. Confocal images were then used to quantify the numbers of cells with stress granules

using Image J software. Results shown are the percentage of cells displaying SGs compared

to the control, and are the mean of three independent experiments (ns: not significant, ** is

P ≤0.01).

160

3.12 Discussion

Caliciviruses, in common with other RNA viruses, utilize the host translational

machinery in order to produce viral proteins. They however use a novel mechanism for

translation as they produce viral RNAs that lack a 5’ cap structure but instead have a virus-

encoded VPg protein covalently linked to the 5’ end of their mRNAs. VPg interacts with

eukaryotic initiation factors (eIF) and acts as a proteinaceous cap-substitute. Previously, we

have demonstrated that feline calicivirus (FCV) VPg interacts with eIF4E to direct translation,

whereas murine norovirus (MNV) VPg interacts with eIF4E and eIF4G, but only the interaction

with eIF4G is important for viral translation (Chaudhry et al., 2006; Chung et al., 2014).

Recently, results from our laboratory demonstrated that calicivirus replication is dependent on

the cell signalling pathways that control the phosphorylation of eIF4E, the MAPK pathway

(Royall et al., 2015). This also suggests that eIF4E phosphorylation is important in regulating

the translation of a subset of host mRNAs implicated in the antiviral response to calicivirus

infection (Royall et al., 2015). In addition to controlling transcription and the activity of the

protein synthesis machinery, recent studies have suggested that the assembly of stress granules

is central in orchestrating stress and antiviral responses to restrict viral replication (Onomoto

et al., 2014). Therefore, using MNV-1 and FCV as models, we investigated whether

caliciviruses control the host response to infection by modulating stress granule responses.

161

3.12.1 Absence of stress granules in stressed and MNV-infected RAW264.7 or BV-2

macrophages.

Using different MNV-permissive cells, including RAW264.7 and BV-2 cells, we could

not detect the assembly of SGs during MNV1 infection (Figure 3.7, 3.14-3.16) and only limited

assembly of granules containing G3BP1, but not TIA-1, and eIF3η, following oxidative stress

with sodium arsenite. These results may reflect a deficiency in RAW264.7 and BV-2 cells,

both of which are macrophages, in their ability to form stress granules. Alternatively, this could

reflect that the antibodies used could be species-specific and not cross-react with other species

such as murine.

It has been shown previously that the overexpression of a protein regulating the

inflammatory response and immune homeostasis, MCPIP1 (monocyte chemotactic protein-

induced protein 1), can impair the assembly of SGs in response to cellular stresses. Whilst

overexpression of MCPIP1 sensitized RAW264.7 cells to stress-induced apoptosis, MCPIP1-

deficient cells are resistant to stress-mediated apoptosis (Qi et al., 2011). MCPIP1 functions

as a deubiquitinase (DUB) to inhibit several signalling pathways such as c-Jun N-terminal

kinase (JNK) signalling pathways (Liang et al., 2010). In addition, MCPIP1 displays RNase

activity towards some inflammatory cytokine mRNAs and viral RNA (Lin et al., 2013;

Mizgalska et al., 2009). As a consequence MCPIP1 inhibits replication of HCV, JEV and

DENV (Lin et al., 2013). However, the ability of MCPIP1 to inhibit SG formation is dependent

on its deubiquitinating enzyme activity and it can impair the assembly of SGs following

oxidative stress with sodium arsenite (Qi et al., 2011). It was suggested that stress results in

the overexpression of MCPIP1 which in turn inhibited arsenite-induced eIF2α phosphorylation.

Therefore, we could hypothesize that the absence of SGs during MNV-1 infection in

this cell type reflects a mechanism in which MNV-1 infection blocks SG formation through

162

the overexpression of MCPIP1, and this could be conserved in BV-2 cells. To fully understand

the role of MNV-1 in SG assembly future work should include the identification of a cell-line

supporting MNV-1 infection and being responsive to oxidative stress, so that positive control

induction experiments are allowed. Alternatively, an MCPIP1-deficient RAW264.7 cell line

could be used for MNV infection and subsequent detection of stress granules, in combination

with monitoring the levels of MCPIP1 by immunoblotting during MNV infection.

3.12.2 FCV replication impairs stress granule assembly during infection

The activation of stress activated kinases by viral infection, mainly PKR, but also

PERK, results in the phosphorylation of eIF2α (Holcik & Sonenberg, 2005). This in turn

prevents the recycling of the ternary complex tRNAiMet-GTP-eIF2, thereby stalling the

initiation of translation resulting in a shutdown of protein synthesis and causing SGs to

accumulate in the cytoplasm (Anderson & Kedersha, 2008). Using G3BP1 and PABP as

markers for SGs, and CRFK and FEA as cells supporting FCV replication, we could not find

evidence of stress granule induction during FCV infection (Figure 3.8). Only uninfected cells

displayed SGs and this observation is dissected further in Chapter 4.

The resulting stoppage in protein synthesis following SG assembly is problematic for

viruses as all viruses rely on the host cell protein synthesis machinery for production of their

proteins. Consequently viruses have evolved many strategies to ensure that viral mRNA

translation can continue by disrupting SG formation during infection or exploiting SGs for their

replication (Montero & Trujillo-Alonso, 2011; Reineke & Lloyd, 2013; Valiente-Echeverria et

al., 2012). The absence of SGs in infected cells during FCV infection could therefore reflect

an inhibition of stress responses during FCV infection.

163

Amongst the different stresses that lead to translational arrest and the formation of SGs,

the most common is dependent on phosphorylation of the α-subunit of eukaryotic initiation

factor 2 (eIF2) at Ser51(Kedersha et al., 1999; McEwen et al., 2005). Typically, sodium

arsenite activates the HRI eIF2α kinase causing oxidative stress-induced phosphorylation of

eIF2α (McEwen et al., 2005). However, translational arrest can also occur independently from

eIF2α phosphorylation through the disruption of the eIF4F complex (Dang et al., 2006; Emara

et al., 2012). Pateamine A (PatA) inhibits eukaryotic translation initiation eIF4A RNA helicase

activity while hydrogen peroxide induces 4EBP hypo-phosphorylation and sequestration of

eIF4E from eIF4F (Emara et al., 2012).

Using sodium arsenite and hydrogen peroxide as stress inducers, representative of the

eIF2α-dependent and independent pathways, respectively, my results showed that FCV

infection impairs SG assembly (Figure 3.17 and 3.18). Only 7% of FCV-infected cells

challenged with sodium arsenite displayed SGs, compared with 83.4% for sodium arsenite-

stressed cells. Likewise 7.7% of FCV-infected cells challenged with hydrogen peroxide

displayed SGs, compared with 87.7% for hydrogen peroxide-stressed cells. Therefore this

would suggest that FCV infection impairs the assembly of stress granules triggered both in an

eIF2α dependent and independent manner. However, because some studies suggested that

H2O2 does not induce SGs (Arimoto et al., 2008; Kedersha & Anderson, 2007), additional

proof of FCV inhibition of eIF2-independent SG assembly could be provided using

Pateamine A or Hippuristanol as inhibitors of eIF4A activity.

The strategies developed by viruses to inhibit stress granules include: (i) proteolytic

degradation of RNA granule factors by viral proteases, (ii) regulation of eIF2α phosphorylation

and (iii) co-opting of RNA granule factors for a new role in viral replication or sequestration

reviewed in (Reineke & Lloyd, 2013). Because FCV infection seems to inhibit both eIF2-

dependent and independent pathways, and given that a previous Ph.D student at Surrey reported

164

eIF2α phosphorylation during FCV infection (Alessandro Natoni, thesis manuscript), it is

likely that SG assembly is inhibited by either (i) proteolytic degradation of RNA granule factors

by viral proteases and (iii) co-opting of RNA granule factors for a new role in viral replication

or sequestration. This would exclude an inhibition mechanism whereby eIF2α phosphorylation

is regulated during infection. These two mechanisms rely on viral products, RNA or proteins

to actively impair SG assembly. Because infection with UV-inactivated virus leads to efficient

SG aggregation following sodium arsenite treatment, this indicates that FCV replication, and

therefore either viral protein(s) or RNA is required to mediate SG inhibition (Figure 3.27).

Several previously identified inhibition mechanisms involve the targeting of the SG-

nucleating protein G3BP1. Non-structural proteins of several alphaviruses interact with G3BP1

in viral complexes that contain nsp3 or nsp2 and nsp4 (Cristea et al., 2010; Fros et al., 2012).

SFV hijacks G3BP1 from SGs and relocates it into viral replication complexes via an

interaction with the C-terminal repeat domains of nsp3, disassembling SGs (Panas et al., 2015;

Panas et al., 2012). As a consequence, SG induction by the eIF2-independent inducer

Pateamine A is blocked (Panas et al., 2012). Similarly, SGs do not form upon infection with

JEV and the JEV core protein directly binds Caprin1 preventing mRNA recognition and SG

formation (Katoh et al., 2013). Viral proteinase from picornaviruses can disperse stress

granules through the cleavage of the SG-nucleating protein G3BP1 by the conserved 3C

proteinases (3Cpro) (Ng et al., 2013; White et al., 2007). Caliciviruses encode a proteinase

designated as a 3C-like proteinase (3CLpro) because of its sequence similarity to the

picornavirus 3Cpro in the active site region (Sosnovtseva et al., 1999), so it is tempting to

speculate that the FCV inhibition of SGs might results from G3BP cleavage. However, using

cell lysates from FCV-infected cells that I generated at the end of my laboratory work,

preliminary immunoblotting experiments suggest that G3BP1 is not cleaved during infection.

However, we cannot exclude that the 3CLpro plays a role in SG disruption by cleaving another

165

SG component. To explore further the role of the FCV 3CLpro, further studies should take

advantage of the possibility of overexpressing a wild-type FCV 3CLpro or an inactive mutant

in CRFK cells (Ian Goodfellow, personal communication). Examination of SG assembly

following sodium arsenite treatment under both conditions should reveal whether the FCV

3CLpro plays any role in the inhibition of SG observed during FCV infection.

Another effective way by which viruses can inhibit stress granule assembly is by

redirecting key SG nucleating proteins into various viral replication foci or other subcellular

localization. For example, IAV can inhibit SG formation by several mechanisms which include

the inactivation of PKR through its binding with non-structural protein 1 (NS1), which in turn

prevents eIF2α phosphorylation and blocks SG formation, while another mechanism requires

the nucleoprotein (NP) without affecting eIF2a phosphorylation (Khaperskyy et al., 2014). In

addition, the host-shutoff protein, namely the polymerase-acidic protein-X (PA-X), can

strongly inhibit SG formation by inducing the concomitant depletion of cytoplasmic poly (A)+

mRNA and the accumulation of poly (A)-binding protein in the nucleus, which impairs the

assembly of PABP-containing SGs later in infection despite eIF2α phosphorylation

(Khaperskyy et al., 2014). Using PABP as SG marker, it was also evidenced that FCV-infection

impairs the assembly of SGs following oxidative stress (Figure 3.10).

Several studies have previously reported that nucleolin, a ubiquitous multifunctional

nucleolar shuttling phosphoprotein, can interact with proteins that are known components of

SGs such as HuR, Staufen, PABP and G3BP1(Isabelle et al., 2012; Tominaga et al., 2011;

Villace et al., 2004; Zheng et al., 2008). Furthermore, nucleolin interacts with the 3’UTR of

NoV and FCV genomic RNA, and the viral RdRp, participating in the replication complex

localized in perinuclear locations (Cancio-Lonches et al., 2011). Therefore, another hypothesis

could be that during infection FCV destabilizes SG assembly by relocalizing SG components

166

to replication complexes. Further studies should aim at dissecting the potential co-localisation

of SG markers within FCV replication complexes using immunofluorescence studies.

Overall the results obtained in this chapter suggest that FCV infection has the ability to

prevent stress responses and avoid the inhibition of protein synthesis, thus enhancing virus

replication. Further studies should be performed to further confirm the impact of FCV infection

on the activation of various virus-related stress pathways previously linked to SGs such as

members of the PKR, RIG-I or c-JUN pathways using immunoblotting.

167

Chapter 4

Paracrine induction of Stress

Granules by

FCV-infected cells

168

4.1 Generation of virus-free cell culture supernatant.

The results from the previous chapter strongly suggest that FCV infection impaired

the ability of CRFK cells to assemble SGs during infection. However, during the infection

SGs could be detected in cells that displayed no sign of infection (revealed by p76 staining)

but resided in the vicinity of infected cells, with 15% of cells displaying SGs at 4h post-

infection. This may reflect a cell-to-cell, paracrine, induction via a chemical or biological

signal of SG formation to dampen viral propagation and/or mount an antiviral response in

uninfected cells.

The only studies that have revealed a mechanism that may be related to this originate

from the observation of mosquito cells infected with Sindbis virus or Semliki Forest virus

(Newton & Dalgarno, 1983; Riedel & Brown, 1979). Cells from the mosquito species Aedes

albopictus, persistently infected with Sindbis virus, produce a low molecular-weight

biomolecule with antiviral activity, capable of specifically affecting Sindbis virus replication.

Unlike interferon, this antiviral agent is virus-specific and cell-specific. Furthermore its

activity is inactivated by treatment with protease K enzyme and heat, but not in response to

anti-Sindbis antibodies (Riedel & Brown, 1979). A similar effect on virus production was

also observed in mosquito cells treated with antiviral material released into the medium from

Semliki Forest virus (SFV)-infected cells. Again this antiviral medium was inactivated by

heat and proteinase K and is specifically associated with alphaviruses and has no significant

effect on other types of arbovirus (Newton & Dalgarno, 1983). However, despite being

identified more than 30 years ago, no mechanistic details have been further elucidated.

169

Therefore, we reasoned that FCV-infected cells may be releasing a signal that would

be able to act as a paracrine inducer of SG formation. To test this hypothesis, we first

generated virus-free supernatant from FCV-infected cells. To this end, CRFK cells were

mock or FCV infected for 6h. The cell culture supernatant was then collected and the viral

particles were UV-inactivated, precipitated using PEG3350 and NaCl and eliminated by

centrifugation. Subsequently, the collected supernatant was assayed for the presence of viral

particles by measuring viral titres using TCID50 assays, and incubating CRFK cells with

supernatants followed by virus detection using immunofluorescence. As shown in Figure 4.1,

viral replication could not be detected by TCID50 following virus elimination as described

above. Thus, we assumed that FCV particles were eliminated from the cell culture

supernatants.

170

FC

V 6

h

FC

V-

su

pern

ata

nt

6h

1 0 0

1 0 1

1 0 2

1 0 3

1 0 4

1 0 5

1 0 6

1 0 7

1 0 8

1 0 9T

CID

50

/mL

Figure 4.1 Detection of FCV replication by TCID50 assay following virus elimination.

CRFK cells were incubated with FCV-infected cell supernatant for 6h, viral titre was then

determined by TCID50 assay. Results show viral titre from supernatant-treated cells compared

to the viral titre from untreated infected cells.

171

4.2 Effect of FCV-infected cell supernatant on Stress granule formation.

Having confirmed that we were able to collect cell culture supernatants free of viral

particles, we proceeded to assay whether these were able to mediate SG formation in

uninfected CRFK cells. To this end, CRFK cells were mock- or FCV-infected for 6h and the

supernatant collected. Viral particles were then eliminated using the procedure described in

section 2.16 (Materials and Methods). The supernatants were then incubated with uninfected

CRFK cells for 1 and 6h. Subsequently the presence of SGs was monitored using

immunofluorescence with G3BP1 as marker, and p76 as marker for viral replication (as a

negative control).

As shown in Figure 4.2, the incubation of CRFK cells with cell supernatant collected

from mock-infected cells did not induce the formation of SGs, reflected by the absence

G3BP1 cytoplasmic foci, either following 1h or 6h of incubation (Figure 4.2, Figure 4.3). In

contrast, the supernatant from FCV-infected cells, was able to induce SG formation in CRFK

cells, with 26.2% of cells displaying SGs after 1h of incubation and 41.7% after 6h.

Furthermore, immunofluorescence microscopy confirmed that no viral particles were present

as reflected by the absence of signal when staining for the viral marker p76 (Figure 4.2).

Therefore these results suggest that FCV-infected cells are able to mediate the release of a

signal that induces SG assembly in neighbouring cells.

172

Figure 4.2 Induction of stress granules by cell supernatant. CRFK cells were incubated with

supernatant from mock or FCV-infected cells for 1h and 6h and then immunostained for G3BP1

antibody (red) and p76 antibody (green) as markers for stress granule and virus respectively,

followed by Alexa-555 and Alexa-488 secondary. Left panels show p76 alone (green), middle

panels show G3BP1 alone (red) and right panels show an overlay of both. Cell nuclei were

stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser

microscope. The bar indicates a size of 10 µm.

Overlay G3BP1 p76

Overlay G3BP1 p76

Overlay G3BP1 p76

Overlay G3BP1 p76

Overlay G3BP1 p76

- sup

+ mock-sup (1h)

+ FCV-sup (1h)

+ mock-sup (6h)

+ FCV-sup (6h)

173

mock-s

up 1

h

FC

V-s

up 1

h

mock-s

up 6

h

FC

V-s

up 6

h

0

2 0

4 0

6 0

8 0

1 0 0

Ce

lls

dis

pla

yin

g

str

es

s g

ran

ule

s (

%)

***

*

Figure 4.3 Quantification of the number of cells displaying stress granules following

treatment with cell supernatant. CRFK cells were incubated with mock or FCV-infected

cell supernatants (sup) and immunostained for G3BP1 as described in Figure 4.1. Image J

software was used to quantify the number of cells displaying SGs. Results show the

percentage of cells with SGs induced by supernatant from FCV-infected cells compared to

either the mock-infected supernatant or untreated cells, and are the mean of three independent

experiments (+/-SE; *** is P≤0.001 for 1h and * is P ≤0.5 for 6h).

174

4.3 Effect of interferon α on stress granule formation in CRFK cells

The production of interferon can be induced in animal cells as the innate immune

response by a variety of antigens and especially by virus infection (Friedman, 1977; Randall

& Goodbourn, 2008). Viral RNA activates signalling pathways that result in the production

of interferons (IFNs) and IFN-stimulated genes which in turn trigger an intracellular antiviral

state that leads to inhibition of a wide range of viruses (Onomoto et al., 2014; Umbach &

Cullen, 2009). Recognition of RNA viruses by TLR7/8 results in the induction of IFN

production via the activation of IRF5 and IRF7 signalling cascades, which substantially leads

to the activation of IFN stimulated genes that mediate the antiviral response (Perry et al.,

2005). Previously it was shown that addition of type I IFN (IFNβ) to cells infected with

measles virus enhanced the assembly of SGs, and IFNβ treatment alone, in the absence of

infection, induced SG formation in ADAR1-deficient cells(John & Samuel, 2014). Moreover

it was reported that IFNβ treatment of HCV-infected cells strongly enhances HCV-induced

phosphorylation of PKR and eIF2α, resulting in increasing SG formation (Garaigorta et al.,

2012; Ruggieri et al., 2012). Based on the general antiviral properties of interferon and these

studies, we reasoned that FCV-infected cells may simply release IFN as a response to

infection, and that this could act as a potent paracrine inducer of SG assembly.

To test this hypothesis, we used commercial feline IFN to investigate the effect of

type I IFN on the accumulation of SGs in CRFK cells. Previous studies have shown that

CRFK cells are responsive to IFN treatment (Baldwin et al., 2004; Harun et al., 2013). CRFK

cells were treated with increasing amount of recombinant feline IFN from 150 IU/ml to

3000 IU/ml and the formation of SGs was monitored by immunofluorescence using G3BP1

as SG marker. As shown in Figure 4.4 and 4.5, the addition of increasing amount of IFNdid

175

not induce any significant increase in the number of SGs, as reflected by the absence of

cytoplasmic G3BP1 foci (Figure 4.4). Therefore, we speculate that type I IFN, at least IFN,

is unlikely to play a role in the paracrine induction of SGs by FCV-infected cells. We

however acknowledge that we did not test directly whether the recombinant IFN used

triggered a response in CRFK cells, however this was published before as mentioned above.

176

0 IU/ml

150 IU/ml

200 IU/ml

300 IU/ml

3000 IU/ml

overlay G3BP1 ToPro-3

overlay G3BP1 ToPro-3

overlay G3BP1 ToPro-3

overlay G3BP1 ToPro-3

overlay G3BP1 ToPro-3

Figure 4.4 Effect of interferon treatment on stress granule formation. CRFK cells were

incubated with increasing concentrations of IFN-α for 6h, then immunostained for G3BP-1

antibody as marker for stress granule. Cell nuclei were stained with ToPro-3 (blue) (left panels)

and middle panels show G3BP alone (red) and right panels show an overlay of both. Cells were

visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10

µm.

177

0150

200

300

3000

0

1 0

2 0

3 0

4 0

5 0

IF N ( IU /m L )

Ce

lls

dis

pla

yin

g

str

es

s g

ran

ule

s (

%)

n s

n s

n s

n s

Figure 4.5 Quantification of the number of cells displaying stress granules following

treatment with IFNα. CRFK cells were incubated with increasing concentrations of IFNα and

immunostained for G3BP1 as described in Figure 4.4. Image J software was used to quantify

the number of cells displaying SGs. Results show the percentage of IFNα-treated cells with SGs

compared to mock-treated cells.

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4.4 Effect of Ribonuclease A (RNase A) on the ability of FCV-infected cell supernatant

to induce stress granule assembly

The accumulation of an RNA replication intermediate during infection leads to the

activation of PKR, which then triggers several downstream effectors involved in the

production of pro-inflammatory cytokines and stress pathways resulting in SG assembly

(Bonnet et al., 2000; Onomoto et al., 2014). Furthermore, in response to infection, host and

viral miRNAs play an important role in regulating expression of viral and host genes

involved in the antiviral response (Emde & Hornstein, 2014; Leung & Sharp, 2010).

Therefore RNA-derived molecules can play key roles in antiviral signalling.

Given that type I IFN is not responsible for the induction of SGs mediated by FCV-

infected cell supernatant, we hypothesise that this as yet unidentified messenger could have

an RNA origin, as various forms of RNA can act as signals for antiviral response. To test this,

we treated the supernatant with RNase A as described in the methods section, a pyrimidine-

specific endonuclease which is commonly used to digest RNA polymers. The ability of the

RNase A-treated supernatant to induce SG assembly in CRFK cells was then monitored using

immunofluorescence microscopy using G3BP1 as SG marker (Figure 4.6). As shown in

Figure 4.6, the digestion of RNA polymers had no effect on the ability of FCV-infected cell

supernatant to induce SG assembly with 25.3% of cells displaying SGs compared with 31%

for untreated supernatant (Figure 4.6, Figure 4.7). Therefore these results suggest that free

RNA polymers are not involved in the signalling from infected to uninfected cells that

triggers SG assembly.

179

Overlay G3BP1 ToPro-3

Overlay G3BP1 ToPro-3

Overlay G3BP1 ToPro-3

Untreated cells

+ FCV-sup/

- RNAse A

+ FCV-sup/

+ RNAse A

Figure 4.6 Effect of RNase A treatment on stress granule formation. FCV-infected cell

supernatants were treated with 7U of RNAse A, and incubated with CRFK cells for 6h, then

immunostained for G3BP1 antibody as marker for stress granule (red). Cell nuclei were stained

with ToPro-3 (blue) (left panels) and middle panels show G3BP1 alone (red) and right panels

show an overlay of both. Cells were visualised using a ZEISS LSM META confocal laser

microscope. The bar indicates a size of 10 µm.

180

FC

V-s

up

FC

V-s

up +

RN

Ase A

Untr

eate

d c

ells

0

1 0

2 0

3 0

4 0

5 0C

ell

s d

isp

lay

ing

str

es

s g

ran

ule

s (

%)

n s

Figure 4.7 Quantification of the number of cells displaying stress granules following

treatment with RNase A. CRFK cells were incubated with 7U RNAse A and

immunostained for G3BP1 as described in Figure 4.6. Image J software was used to

quantify the number of cells displaying SGs. Results show the percentage of cells with

SGs induced by supernatant (sup) from FCV-infected cells compared with cells treated

with FCV-cell supernatant plus RNase, and are the means of three independent

experiments (+/-SE; is ns: not significant).

181

4.5 Effect of Heat-Shock on the ability of FCV-infected cell supernatant to induce stress

granule assembly

The infection of cells by viruses is a trigger for a large array of innate immune

responses. Several immune receptors expressed on the surface of cells or internally can ignite

that immune response, the major one being Toll-like receptors (TLRs), retinoic-acid

inducible gene I (RIG-I) and NOD-like receptors (NLRs). RNA viruses can activate the TLRs

displayed on endosomal compartments such as TLR3, TLR7, TLR8 and TLR9 (Pichlmair &

Reis e Sousa, 2007), while replication intermediates also activate the PKR pathway (Garcia et

al., 2006). These events result in the production of many pro-inflammatory cytokines and

chemokines (such as TNF, IL-1 or IL-8) and IFN that are secreted and can amplify the

immune response via both autocrine and paracrine effects, and further impact on the outcome

of infection (Rouse & Sehrawat, 2010).

Therefore we investigated whether the induction of SGs by virus-free supernatants

could be mediated by the release of protein messengers, putatively an immune mediator of a

proteinaceous nature. To test this hypothesis, we performed heat-shock of the supernatant to

denature proteins by heating supernatants at 65°C for 10 or 20 min. The ability of the heat-

shocked supernatant to induce SG assembly in CRFK cells was then monitored using

immunofluorescence and G3BP1 as SG marker (Figure 4.8). As shown in Figure 4.8, the

denaturation of proteins impaired the ability of FCV-infected cell supernatant to induce the

formation of SGs. This was reflected by a reduction in G3BP1 cytoplasmic foci following 10

min inactivation, dropping from 30.9% to 0.5%, and a complete disappearance of SGs

following 20 min of inactivation (Figure 4.8, Figure 4.9). Therefore these results could

182

suggest that the signalling from FCV-infected to uninfected cells involves a messenger of

proteinaceous nature.

183

Untreated cells

FCV-sup

FCV-sup + 10 min heat

shock

FCV-sup + 20 min heat

shock

Figure 4.8 Effect of heat shock on stress granule formation. FCV-infected cell

supernatants (FCV-sup) were exposed to heat shock for 10 or 20 min, incubated with

CRFK cells for 6h, then immunostained for G3BP1 antibody as markers for stress

granule (red) followed by Alexa-555 secondary antibody. Cell nuclei were stained

with ToPro-3 (blue) (left panels) and middle panels show G3BP alone (red) and right

panels show an overlay of both. Cells were visualised using a ZEISS LSM META

confocal laser microscope. The bar indicates a size of 10 µm.

Overlay G3BP1 ToPro-3

Overlay G3BP1 ToPro-3

Overlay G3BP1 ToPro-3

Overlay G3BP1 ToPro-3

184

FC

V-s

up

FC

V-s

up

+ H

eat-

sh

ock 1

0 m

in

FC

V-

su

p+ H

eat-

sh

ock 2

0 m

in

Un

treate

d c

ells

0

2 0

4 0

6 0

8 0

1 0 0C

ell

s d

isp

lay

ing

str

es

s g

ra

nu

les

(%

)

* * *

* * *

n s

Figure 4.9 Quantification of the number of cells displaying stress granules following

treatment with heat-shocked supernatant. Confocal microscopy was used to detect the

effect of heat shock on stress granule formation as described in Figure 4.8, Image J software

was then used to quantify the number of cells displaying SGs following treatment with heat-

shocked supernatant. Results show the percentage of cells with SGs induced by heat-shocked

FCV-infected cell supernatant compared with cells treated with unheated FCV-infected cell

supernatant, and are the means of three independent experiments (+/-SE; *** is P ≤0.001, ns

not significant).

185

4.6 Effect of FCV-infected cell supernatant on a wide range of cells’ ability to assemble

stress granules

Having now proposed that FCV-infected CRFK cells release a proteinaceous

messenger that can mediate the assembly of SGs, we next ask whether this messenger could

have a ubiquitous action and induce the assembly of SGs in other cell lines. To test this

hypothesis, we incubated the virus-free supernatant of FCV-infected CRFK cells with

different cells, including FEA (feline cells supporting FCV replication), 293T (a human

embryonic kidney cell line supporting FCV replication), RAW264.7 (murine macrophage

cells supporting MNV-1 replication), BV-2 (murine microglia cells supporting MNV-1

replication) and J774 (murine macrophage cells supporting MNV-1 replication) and THP-1

(human macrophage cells), for 6h. The presence of SGs was then monitored by

immunofluorescence using G3BP1 as marker. G3BP1 cytoplasmic foci, representative of

SGs, were clearly detected in FEA and 293T cells with 71.6% and 57.3% of cells displaying

SGs, respectively (Figure 10, Figure 11). In stark contrast, no SG induction could be detected

in RAW264.7 and BV-2 cells (Figure 10, Figure 11) as well as THP-1 and J774 (data not

shown). Of note these cell lines are all murine or human macrophage cell lines. Therefore,

although the supernatant of FCV-infected CRFK cells has the ability to induce SG formation

in other cell types, it does seem to exhibit a level of cell specificity, and could be related to

the fact that these cells support FCV replication.

186

Figure 4.10 Effect of FCV-infected cell supernatant on a wide range of cells. 293T, FEA, RAW264.7, BV-2,

THP-1 and J774 cells were grown as described in Materials and Methods, then treated with supernatant from

FCV-infected cells (FCV-sup) for 6h. Untreated and treated cells were then immunostained for G3BP1 (red) and

p76 (green) as markers for stress granule and virus respectively, followed by Alexa-555 and Alexa-488

secondary antibodies. Left panels show untreated cells (-FCV-sup), right panels show treated cells (+FCV-sup).

Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a ZEISS LSM META confocal laser

microscope. The bar indicates a size of 10 µm.

G3BP1 G3BP1

G3BP1 G3BP1

THP-1

J774

G3BP1 G3BP1

293T

- FCV-sup + FCV-sup

G3BP1

FEA

G3BP1

G3BP1 G3BP1

RAW264.7

BV-2

G3BP1 G3BP1

187

293 c

ells

FE

A c

ells

RA

W 2

64.7

cells

BV

-2 c

ells

TH

P-1

cells

J774 c

ells

0

2 0

4 0

6 0

8 0

1 0 0

Ce

lls

dis

pla

yin

g

str

es

s g

ra

nu

les

(%

)

U n tre a te d c e lls

F C V -s u p

Figure 4.11 Quantification of the number of cells displaying stress granules following

treatment of a wide range of cells with FCV-infected cell supernatant. CRFK cells

were incubated with mock- or FCV-infected cell supernatants and immunostained for

G3BP1 as described in Figure 4.10. Image J software was used to quantify the number of

cells displaying SGs. Results show the percentage of cells with SGs induced by

supernatant from FCV-infected cells compared with untreated cells.

188

4.7 Discussion

4.7.1 FCV infection leads to paracrine induction of SG assembly

Because of the ability of viruses to trigger the formation of SGs and cause

translational shut-off, it has been proposed that SGs could exert a specific antiviral effect,

both on viral mRNA and/or by regulating mRNA contributing to the antiviral response

(Onomoto et al., 2014). Following our observation that SGs can be detected in some

uninfected cells during FCV infection, we hypothesize that FCV-infected cells may release a

signalling molecule that could trigger SG assembly in neighbouring cells, which would be

consistent with this proposed antiviral role for SGs.

To test this hypothesis, I generated virus-free supernatants from mock- and FCV-

infected cells. These supernatants were first tested to confirm the absence of viral replication

by determining viral titre using a TCID50 assay (Figure 4.1) as well as an

immunofluorescence assay to detect viral protein (Figure 4.2). Neither method detected viral

protein or viral replication confirming that the supernatants were indeed virus-free.

The supernatants were then added to CRFK cells and the presence of SGs was

examined by immunofluorescence microscopy (Figure 4.2). While the supernatant from

mock-infected cells did not trigger SG formation, 26% of cells treated with virus-free

supernatant from FCV-infected cells formed SGs after 1h and 42% after 6h (Figure 4.3).

These results suggest that FCV-infected cells release a soluble mediator that induces SG

assembly in neighbouring cells. This paracrine induction of SGs in uninfected cells has never

been described and may reflect a novel regulatory mechanism to control stress during

infection by modulating mRNA metabolism.

189

4.7.2 Different types of stress granules can be assembled in response to infection

Recent studies have suggested that SGs have an antiviral role and function as a

cellular platform to initiate innate immune responses (Onomoto et al., 2014). SGs can serve

as sentinel mechanisms for IFN activation by promoting the condensation of IFN signalling

moieties with their viral RNA ligands (Khaperskyy et al., 2014; Langereis et al., 2013;

Onomoto et al., 2012; Yoo et al., 2014). In addition to canonical SG markers, these antiviral

SGs contain viral RNA and nucleoproteins. Thus, stress granules may be important in

priming neighbouring cells against infection by (i) suppressing viral replication through

inhibition of viral protein synthesis, and (ii) serving as a platform to facilitate IFN production.

In addition to these, viruses have also been linked to the formation of antiviral granules

(AVGs) (Rozelle et al., 2014). The disruption of Poxvirus replication triggers the formation

of host-protein dense antiviral granules (AVGs) which differ from SGs by their localization

to cytoplasmic viral factories and by their composition (Rozelle et al., 2014). While these

AVGs share many components with SGs, they are characterised by the absence of eIF3 and

40S subunits, and their stability in the presence of translation elongation inhibitors. This

suggested that AVGs may not act as sorting sites for mRNAs but exists to restrict viral

translation. Overall a link between antiviral immunity and stress granules is clear, there seem

to be different specific responses linked to different types of RNA granules.

To investigate the nature of the stress granules induced by virus-free supernatant,

further studies should aim at investigating whether these are dissolved following

cycloheximide treatment, as canonical SGs. Then, the composition of these SGs should be

characterized using immunofluorescence microscopy to detect a variety of canonical SG

markers (G3BP1, Caprin, TIA1, PABP, eIF4G), IFN signalling factors previously found to

accumulate in avSGs (RIG-I, MDA5, PKR) and markers that accumulate in SGs but not

AVGs (40S, eIF3 and poly(A)+ mRNA using digoxigenin-labeled oligo-d(T) and anti-

190

digoxigenin antibody). Furthermore, because canonical SG assembly mostly results from

translational arrest, further work should aim at investigating whether the assembly of SGs

mediated by the supernatant of infected cells is the consequence of a translational shut-off.

To determine whether translation is inhibited in the subset of cells that form SGs,

visualization of de novo protein synthesis by metabolic incorporation of the amino acid

analog L-azidohomoalanine (AHA) could be employed. After cell fixation, proteins that had

incorporated AHA can be conjugated with a fluorescent probe, such as Alexa Fluor 594,

using a copper-catalysed reaction and labelling any nascent peptide with AHA-594 as

described previously (Rozelle et al., 2014). As a control, cells should be treated with the

translation inhibitor cycloheximide, which should completely block AHA-594 staining. Cells

would also be stained for stress granule markers, such as G3BP1, to score both stress granule

formation and active translation in cells. This would identify any link between inactive

translation and SG induction, providing evidence as to whether virus-free supernatant

mediates a translational shut-off.

4.7.3 Could this paracrine induction of SGs reflect an antiviral mechanism?

Previously, some studies have described the release of a low molecular weight

antiviral agent into the medium of A.albopictus cells persistently infected with Sindbis virus

(Riedel & Brown, 1979), or Semliki Forest virus (Newton & Dalgarno, 1983). This agent was

able to reduce the normal viral yield. Treatment of mosquito cultures with the supernatant

containing the antiviral agent before infection prevents the normal production of high virus

yield (Newton & Dalgarno, 1983; Riedel & Brown, 1979). However, the antiviral activity

was sensitive to proteinase K and heat, being rapidly inactivated at 56˚C. These studies

reported that the antiviral agent was also cell and virus specific. While this antiviral activity

191

has been shown to selectively affect Sindbis virus production in mosquito cells, it had no

effect on virus production in BHK-21 cells (Condreay et al., 1988; Condreay & Brown, 1988;

Riedel & Brown, 1979). In addition, Kanthong et al., 2010 discovered that cell-free

supernatant solutions from C6/36 mosquito cell cultures are capable of destabilizing

persistently-infected cultures with Dengue virus. The authors proposed that the cell-free

supernatant contains small polypeptides with cytokine-like activity resulting in an effect on

virus yield and to induce a protective response against DENV-2 virus infection in naïve cells

(Kanthong et al., 2010).

In agreement with these data, our results report that the ability of virus-free

supernatant to induce SGs was sensitive to heat treatment at 65˚C (Figure 4.8). In addition,

our results demonstrated that the effect of this supernatant was not due to RNA, as RNase

treatment had no impact (Figure 4.7). A subsequent experiment was carried out to determine

the nature of this antiviral agent contained within the supernatant. The fact that virus infected

cells could induce an antiviral state in surrounding host cells during most viral infections, led

us to the hypothesis that the induction of SGs by supernatant may be due to interferon

activity. However, treating CRFK cells with feline IFN did not induce SG assembly, showing

that release of IFN from infected cells is unlikely to be responsible for the paracrine SG

induction (Figure 4.4).

To further investigate whether the assembly of SGs in uninfected cells reflects the

induction of a protective antiviral effect, CRFK cells could be treated with virus-free

supernatant from mock or FCV-infected cells and then challenged with FCV. Viral

replication would then be quantified by measuring viral titre using TCID50 assay. The

antiviral state of the cells would then be characterized further by measuring IFN levels using

qRT-PCR. In addition the conservation of this mechanism could be investigated by analysing

192

the ability of virus-free supernatant from cells infected with other viruses to induce SGs,

using model arboviruses and herpesviruses studied at Surrey.

4.7.4 Investigating the nature of the soluble mediator(s) triggering stress granule

assembly

To date, the only example of a paracrine inducer of SG assembly is angiogenin, an

angiogenic factor released by tumour cells (Yamasaki et al., 2009). Angiogenin is a secreted

stress-activated ribonuclease producing tRNA-derived stress-induced RNA (tiRNAs) to

promote translational repression and induce SG assembly (Emara et al., 2010). To investigate

whether angiogenin plays a role in the paracrine induction of SGs in bystander cells, extracts

prepared from CRFK cells treated with virus-free supernatants of mock-infected or FCV-

infected cells could be separated on a denaturing gel and analyzed to visualize stress-induced

small RNA by northern blotting with cDNA probes complementary to the 5’ and 3’ end of

tRNA. In addition, CRFK cells could be transfected with control or Dharmacon SMART pool

siRNA targeting angiogenin to reduce its expression before investigating the effect on virus-

free supernatant ability to induce SGs.

We cannot however rule out that the paracrine inducer of SG assembly could have a

different nature: nucleic acid, such as RNA fragments of viral origin, cellular miRNAs, other

proteins such as cytokines or signalling proteins, or more complex structures such as

exosome particles which can deliver antiviral sensors to uninfected cells (Kalamvoki et al.,

2014). It has been proposed that exosomes may be responsible for transferring anti-viral

signal between cells (Li et al., 2013; Schorey & Bhatnagar, 2008). Exosomes can carry RNA

such as mRNA, miRNA and genomic fragments, and a large component of proteins and

lipids among host cells leads to triggering of an anti-viral state in uninfected cells (Huang et

193

al., 2013; Schorey & Bhatnagar, 2008; Valadi et al., 2007). For example, HIV transfer of the

antiviral cytidine deaminase APOBEC3G via exosomes inhibits HIV replication in recipient

cells (Khatua et al., 2009).

Therefore virus-free supernatant from mock- or FCV-infected cells could be treated

with proteinase K and the ability to induce SG assembly monitored by immunofluorescence

microscopy to confirm whether a protein mediator is involved in the paracrine induction of

SGs. The presence of exosome particles in the supernatant should also be evaluated using

commercial exosome purification procedures and immunoblotting against the exosome

marker CD9 (Zeringer et al., 2015). Proteomics analysis from cell culture supernatant is

difficult due to the presence of highly abundant serum proteins. However supernatant could

be fractionated over a gel filtration column into several fractions, and the individual fractions

could be tested to determine whether they were able to induce SGs. These fractions could

then be analysed by SDS-PAGE and liquid chromatography–mass spectrometry (LC-MS).

Currently, this option is being investigated in collaboration with the proteomics facility at the

University of Cambridge (collaboration with Dr Kathryn Lilley).

Overall, the results from Chapter 4 suggest that a new mechanism of paracrine

signalling during viral infection is involved in the assembly of SGs in uninfected cells and it

is proposed that this unprecedented mechanism could play a key part in the antiviral

response.

194

Chapter 5

Regulation of P-Bodies formation

during

FCV and MNV-1 infection

195

5.1 Optimisation of the markers and conditions used for P-bodies detection

The previous results revealed that the assembly of SGs is impaired during FCV

infection. This prompted us to ask whether caliciviruses also have the ability to modulate the

formation of other cytoplasmic RNA granules in infected cells, namely PBs. To this end,

FCV and MNV-1 were used as surrogate model for human norovirus, in order to examine

their effect on PBs formation.

First, this work began with the optimisation of experimental conditions for

immunofluorescence detection of PB assembly. While PBs are constitutively form in the

cytoplasm of cells under normal conditions, their numbers and size increase upon stress

condition (Teixeira et al., 2005). Therefore, PBs markers optimisation was tested in

unstressed cells that support FCV or MNV-1 replication, CRFK and RAW264.7 cells

respectively, and monitored by immunofluorescence microscopy using Dcp1 and Xrn1 as

markers.

Among different concentrations of Dcp1 antibody tested, cytoplasmic foci were only

detected in unstressed CRFK and RAW264.7 cells at dilution of 1:50, while lower dilution

did not lead to the detection of PBs (Figure 5.1, 5.2 and data not shown). By contrast, no

signal could be detected for Xrn1, even when testing a wide range of dilutions from 1:50 to

1:500 (Figure 5.1, 5.2 and data not shown) using both CRFK and RAW264.7 cells.

196

Figure 5.1 Optimization of Dcp1 and Xrn1 antibody dilution in CRFK cells for

immunofluorescence studies. CRFK cells were fixed and stained for different

concentrations of Dcp1 (A-C) (red) and Xrn1 antibodies (D) antibodies as indicated,

followed by Alexa-555 secondary antibody. ToPro3 stained nuclei are shown in blue. Cells

were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a

size of 10 µm.

ToPro-3 Dcp1 Overlay

ToPro-3 Dcp1 Overlay

ToPro-3 Dcp1 Overlay

A

B

C

1:50 1:100 1:500

D 1:50

Xrn1 Overlay ToPro-3

197

Figure 5.2 Optimization of Dcp1 and Xrn1 antibody dilution in RAW264.7 cells for

immunofluorescence studies. RAW264.7 cells were fixed and stained for different

concentrations of Dcp1 (A) and Xrn1 (B and C) antibodies, as indicated, followed by

Alexa-555 secondary antibodies. ToPro3 stained nuclei are shown in blue. Cells were

visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size

of 10 µm.

1:50

1:100

Xrn1 Overlay ToPro-3

Xrn1 Overlay ToPro-3

B

C

ToPro-3 Dcp1 Overlay

1:50 A

198

As discussed for SGs in section 3.6, the concentration of foetal bovine serum (FBS)

can act as a source of stress on cells and affect the condition for infection. Therefore the

effect of FBS concentration on PB assembly was analysed by incubating CRFK cells with

different concentrations of FBS (0, 2 and 10%) and monitoring the impact on PBs by

immunofluorescence microscopy using Dcp1 as marker (Figure 5.3). Immunofluorescence

analysis revealed no significant differences in numbers of cells displaying PBs or the number

of PBs per cell using these different concentrations (Figure 5.3, and data not show).

Therefore 2% of FBS was chosen for all subsequent studies as we did with SGs.

199

Figure 5.3 Optimization of foetal bovine serum concentration for detect of P-bodies

in CRFK cells. CRFK cells were grown overnight using different concentrations of FBS

(A-C), cells were then fixed and stained for Dcp1 antibody (red; middle panels), followed

by Alexa-555 secondary antibody. ToPro3 stained nuclei are shown in blue. Cells were

visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a size

of 10 µm.

0% FBS 2% FBS 10% FBS

ToPoro-3 Dcp1 Overlay

ToPoro-3 Dcp1 Overlay

ToPoro-3 Dcp1 Overlay

A

B

C

200

5.2 Effect of MNV-1 infection on P-bodies formation.

To investigate whether MNV-1 infection interfere with PBs formation, RAW264.7

cells were infected with MNV-1 at MOI of 30 for 18h, and examined by immunofluorescence

microscopy at 0, 4, 8, 12 and 18h post infection using Dcp1 as a marker for PBs, and NS7 as

marker for MNV-1 (Figure 5.4). Overall, a significant reduction in number of cells displaying

PBs was observed from 8 hpi onwards during MNV-1 infection relative to mock-infected

cells (Figure 5.4). This disruption was reflected with a drop in the number of cells displaying

PBs from 90.4 to 39.4% when compared to mock-infected cells at 8 hpi, 68.6 to 19.2% at 12

hpi, and 99.6 to 28.5% at 18 hpi (Figure 5.5). In summary these experiments suggest that

MNV-1 infection disrupt PB assembly during infection, and that this starts in the middle of

the replication cycle at a time where viral RNA synthesis begins. .

201

0 hpi

4 hpi

8 hpi

- MNV-1 + MNV-1

NS7 DCP1 Overlay

NS7 DCP1 Overlay

NS7 DCP1 Overlay

NS7 DCP1 Overlay

NS7 DCP1 Overlay

NS7 DCP1 Overlay

- MNV-1 + MNV-1

- MNV-1 + MNV-1

202

Figure 5.4 Detection of P-bodies during MNV-1 infection. RAW264.7 cells were infected

with MNV-1 at MOI of 30 for several time points as indicated, and then immunostained for

Dcp1 antibody (red) and NS7 (green) as markers for PBs and virus respectively. Left panels

show NS7 alone (green), middle panels show Dcp1 alone (red) and right panels show an

overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using a

ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.

12 hpi

18 hpi

NS7 Dcp1 Overlay

NS7 Dcp1 Overlay

NS7 Dcp1 Overlay

NS7 Dcp1 Overlay

- MNV-1 + MNV-1

- MNV-1 + MNV-1

203

0 4 812

18

0

2 0

4 0

6 0

8 0

1 0 0

h o u rs p o s t- in fe c t io n

Ce

lls

dis

pla

yin

g

PB

s

(%)

- M N V -1

+ M N V -1n s

****

****

****

n s

Figure 5.5 Quantification of the number of cells displaying P-bodies during MNV-1

infection. RAW264.7 cells were infected with MNV-1 at MOI of 30 for several time points,

stained for Dcp1 and NS7 as described in Figure 5.4. Confocal images were then used to

quantify the numbers of cells with PBs using Image J software. Results shown are the

percentage of cells displaying PBs at each time point compared to the control, and are the

mean of three independent experiments (+/-SE; **** is P ≤0.0001; ns: not significant).

204

To confirm that this effect was not cell-line specific, BV-2 cells were infected with

MNV-1 at MOI of 30 for 12h and the effect on PB assembly was investigated using Dcp1 as

marker for PBs and NS7 as marker for the virus. Similarly to results obtained with

RAW264.7 cells, the infection with MNV-1 also resulted in the reduction of PBs in BV-2

cells (Figure 5.6). As shown on Figure 5.7, 27.8% of MNV-1 infected cells displayed PBs,

against 76.9.6% for mock-infected cells (Figure 5.7).

205

Figure 5.6 Detection of P-bodies during MNV-1 infection using BV-2 cells. BV-2 cells

were infected with MNV-1 at MOI of 30 for 12h, and then immunostained for Dcp1

antibody (red) and NS7 antibody (green) as markers for PBs and virus respectively. Left

panels show NS7 alone (green), middle panels show Dcp1 alone (red) and right panels show

an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells were visualised using

a ZEISS LSM META confocal laser microscope. The bar indicates a size of 10 µm.

NS7 Dcp1 Overlay

NS7 Dcp1 Overlay

- MNV-1 + MNV-1

206

0

2 0

4 0

6 0

8 0

1 0 0

1 2 h o u rs p o s t- in fe c t io n

Ce

lls

dis

pla

yin

g

PB

s

(%)

- M N V -1

+ M N V -1

*

Figure 5.7 Quantification of the number of cells displaying P-bodies during MNV-1

infection in BV-2 cells. BV-2 cells were infected with MNV-1 at MOI of 30 for several

time points, stained for Dcp1 and NS7 antibodies as described in Figure 5.6. Confocal

images were then used to quantify the numbers of cells with PBs using Image J software.

Results shown are the percentage of cells displaying PBs compared to the control, and are

the mean of three independent experiments (+/-SE; * is P ≤0.5).

207

The section 5.1 revealed that using Xrn1 as marker did not allow the detection of PBs

in RAW264.7. However, we cannot exclude that this marker could accumulate in the PBs

formed in RAW264.7 cells during MNV-1 infection. To test this, RAW264.7 cells were

infected with MNV-1 at MOI of 30 for 24h and the assembly of PBs monitored by

immunofluorescence microscopy using Xrn1 and NS7 as marker for PBs and infection

respectively. As shown on Figure 5.8, no cytoplasmic foci could be detected using this

marker at any point during infection. These results confirm that Xrn1 is not a suitable marker

for the detection of PBs either during MNV-1 infection of RAW264.7 cells or during normal

conditions (Figure 5.2, Figure 5.8).

208

0hpi

4hpi

- MNV-1 + MNV-1

8hpi

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

- MNV-1 + MNV-1

- MNV-1 + MNV-1

209

Figure 5.8 Detection of P-bodies during MNV-1 infection using Xrn1 as a marker.

RAW264.7 cells were infected with MNV-1 at MOI of 30 for different time points, and then

immunostained for Xrn1 antibody (red) and NS7 (green) as markers for PBs and virus

respectively. Left panels show NS7 alone (green), middle panels show Xrn1 alone (red) and

right panels show an overlay of both. Cell nuclei were stained with ToPro-3 (blue). Cells

were visualised using a ZEISS LSM META confocal laser microscope. The bar indicates a

size of 10 µm.

12hpi

18hpi

24hpi

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

NS7 Xrn1 Overlay

- MNV-1 + MNV-1

- MNV-1 + MNV-1

- MNV-1 + MNV-1

210

5.3 Effect of FCV infection on the assembly of P-bodies.

To investigate the effect of FCV infection on the formation of PBs, CRFK cells were

infected with FCV Urbana at MOI 0.2 for 6h, and the presence of PBs was investigated by

immunofluorescence microscopy using Dcp1 as marker for PBs and p76 as marker for viral

particles (Figure 5.9). As shown on figure 5.9, FCV resulted in a reduction in PBs abundance.

The quantification of the number of cells displaying PBs revealed that 91% of mock-infected

contained PBs, while only 19% of FCV-infected cells contained PBs (Figure 5.10). These

results suggest that FCV infection, like MNV-1 infection, disrupts PB assembly.

211

Figure 5.9 Detection of P-bodies during FCV infection. CRFK cells were infected with FCV at

MOI of 0.2 for 6h, and then immunostained for Dcp1 antibody (red) and p76 antibody (green) as

markers for PBs and virus respectively. Left panels show p76 alone (green), middle panels show

Dcp1 alone (red) and right panels show an overlay of both. Cell nuclei were stained with ToPro-3

(blue). Cells were visualised using a ZEISS LSM META confocal laser microscope. The bar

indicates a size of 10 µm.

p76 Dcp1 Overlay

p76 Dcp1 Overlay

-FCV

+FCV

212

0

2 0

4 0

6 0

8 0

1 0 0

Ce

lls

dis

pla

yin

g

PB

s

(%)

- F C V

+ F C V 6 h

***

Figure 5.10 Quantification of the number of cells displaying P-bodies during FCV

infection. CRFK cells were infected with FCV at MOI of 0.2 for 6h, stained for Dcp1 and

p76 as described in Figure 5.9. Confocal images were then used to quantify the numbers of

cells with PBs using Image J software. Results show are the percentage of cells displaying

SGs compared to the control, and are the mean of three independent experiments (+/-SE;

*** is P ≤0. 001).

213

5.4 Effect of chemical induction of oxidative stress on P-body assembly

PBs formation is induced by a wide range of experimental stresses such as glucose

deprivation, osmotic stress, and decreased translation initiation rates (Buchan & Parker, 2009;

Iwaki & Izawa, 2012; Teixeira et al., 2005). Importantly, most of these stresses are also

inducing the assembly of SGs (Anderson & Kedersha, 2006; Kedersha & Anderson, 2009;

Kedersha et al., 2005). By contrast, some inducer, such as Clotrimazol, can induce SGs but

not PBs (Buchan et al., 2011), and conversely, acidic stress only induces PBs formation

(Iwaki and Izawa, 2012). In addition, some studies have suggested that the assembly of stress

granules is dependent on the existence of PBs, implying that stress that lead to PB formation

could also induce SGs formation (Buchan et al., 2008). In contrast, other studies support

models in which mRNA exists in equilibrium between polysomes, SGs and PBs (Anderson &

Kedersha, 2009a; Decker & Parker, 2012; Kedersha et al., 2000; Lloyd, 2012) and (Figure

1.4). Finally, the size and numbers of PBs are dependent on the type of stress (Buchan &

Parker, 2009; Teixeira et al., 2005). As such, the induction of PBs by oxidative stress, such as

sodium arsenite or hydrogen peroxide, and the link with SG assembly, is a matter of debate.

Some studies demonstrated that PBs numbers and size increased upon sodium arsenite

treatment (Eulalio et al., 2007a; Eulalio et al., 2007b; Kedersha & Anderson, 2007; Serman

et al., 2007; Shah et al., 2013; Souquere et al., 2009; Wang et al., 2013; Wilczynska et al.,

2005). However, other studies demonstrated that hydrogen peroxide treatment results in the

inhibition of PBs, directly (Shah et al., 2013) or through the inhibition of factors, such as

microtubules, required for PB assembly, such as microtubules (Hu & Lu, 2014; Lee et al.,

2005; Lindsay & McCaffrey, 2011). Therefore, building on the previous results in section 3.2

214

and 3.8, suggesting that CRFK cells are able to form SGs in response to sodium arsenite or

hydrogen peroxide, the impact of these treatments on the assembly of PBs was investigated.

The ability of CRFK cells to respond to stresses that induce SGs assembly was

investigated by treating CRFK cells with 0.5 mM of sodium arsenite for 1h or with 1mM

H2O2 for 2h and subsequently monitoring the assembly of PBs by immunofluorescence

microscopy using Dcp1 as marker (Figure 5.11). Upon hydrogen peroxide treatment, the

number of PBs dropped from 95.2% to 8.8% (Figure 5.12). By contrast, a more modest yet

significant dispersion of PBs was observed upon treatment of cells with sodium arsenite with

75.2% of cell displaying PBs against 90.8% in mock-treated cells (Figure 5.12). These results

suggest that the ability of CRFK cells to form PBs is reduced in response to hydrogen

peroxide and sodium arsenite. Although only preliminary, this could reflect that the formation

of SGs is not strictly dependent on pre-existing PBs in CRFK cells. This would rather support

a model in which mRNAs are in equilibrium between RNA granules and whereby stimulating

the assembly of a type of granules, in this case SGs, conversely has a negative impact on the

abundance of other types, in this case PBs.

215

ToPro-3 Dcp1 Overlay

ToPro-3 Dcp1 Overlay

ToPro-3 Dcp1 Overlay

ToPro-3 Dcp1 Overlay

- H2O2

+ H2O2

- SA

+SA

Figure 5.11 Effect of the sodium arsenite and hydrogen peroxide treatment on P-

body assembly. CRFK cells were treated with 0.5mM sodium arsenite (SA) for 1h or 1

mM of hydrogen peroxide (H2O2) for 2h, stained for Dcp1 as a marker for PBs (red),

followed by Alexa-555 secondary antibody. Cell nuclei were stained with ToPro-3 (blue)

(left panels) and middle panels show Dcp1 alone (red) and right panels show an overlay of

both.

216

- S

A

+S

A

- H

2O

2

+H

2O

2

0

2 0

4 0

6 0

8 0

1 0 0

1 2 0

Ce

lls

dis

pla

yin

g

PB

s

(%)

*****

Figure 5.12 Quantification of the number of cells displaying P-bodies upon treatment

with the sodium arsenite and hydrogen peroxide. CRFK cells were treated with 0.5 mM

of sodium arsenite (SA) for 1h or 1mM hydrogen peroxide (H2O2) for 2h, stained for

Dcp1 as a marker for PBs, followed by Alexa-555 secondary antibody as describe in

figure 5.11. Confocal images were then used to quantify the numbers of cells with P-

bodies using Image J software. Results show the percentage of cells with PBs upon SA or

H2O2 compared to untreated cells, and are the mean of three independent experiments (+/-

SE; * and **** is p ≤ 0.1 and 0.0001 for SA and H2O2 respectively.

217

5.5 Discussion

5.5.1 Modulation of P-bodies assembly during MNV-1 and FCV infection

The translation and degradation of cytoplasmic mRNA is one of several mechanisms

used by eukaryotic cells in order to regulate gene expression (Cougot et al., 2004; Teixeira et

al., 2005). These mechanisms are thought to be competing pathways as the inhibition of

translation could result in an increased mRNA degradation rate (Franks & Lykke-Andersen,

2008). PBs play a role in the regulation of mRNA turnover and contain translationally

repressed mRNAs together with several proteins involved in mRNA decapping,

deadenylation, miRNA mediated mRNA silencing, and mRNA storage (Anderson &

Kedersha, 2009a; Cougot et al., 2004; Eulalio et al., 2007a; Eulalio et al., 2007b; Kedersha &

Anderson, 2009; Kedersha et al., 2005; Parker & Sheth, 2007; Teixeira et al., 2005). They are

found in cells under normal conditions and are conserved among eukaryotes (Decker &

Parker, 2012).

The results from chapter 3 showed that FCV infection impairs the ability of cells to

form SGs in response to different types of stresses. This could suggest that FCV infection has

the ability to prevent the inhibition of cellular protein synthesis, thereby enhancing viral

replication. While previous studies have identified that pre-existing PBs are require for SG

formation (Buchan et al., 2008; Kedersha et al., 2005), other did not (Mollet et al., 2008; Ohn

et al., 2008; Shah et al., 2013). It has also been suggested that the mRNA are in dynamic

equilibrium between polysomes, SGs and PBs (Balagopal & Parker, 2009; Buchan & Parker,

2009). These led to the hypothesis that caliciviruses may also interfere with cytoplasmic

RNA granules at the level of the PBs.

Thus to understand better the relationship between calicivirus infection and

cytoplasmic RNA granules, I investigated the regulation of PBs assembly during FCV and

218

MNV-1 infection. During MNV-1 infection, from 8 hpi, and FCV infection, from 6 hpi, PB

assembly is disrupted, as reflected by a reduction in Dcp1 cytoplasmic foci. MNV-1 infection

resulted in a reduction from 68.6 to 19.2% in the number of cells containing PBs, while FCV

infection resulted in a reduction from 90.8 to 19.1%. These results indicate that MNV-1 and

FCV have the ability to disrupt PBs, an event that may be required for a viral replication.

Viruses have evolved many strategies to counteract PB assembly and evade RNA

decay pathways. This disruption may be due to the modification or cleavage of a specific PB-

nucleating protein or may result from redistribution of PB components to enhance viral

replication (Lloyd, 2013; Reineke & Lloyd, 2013; Tsai & Lloyd, 2014). All these

mechanisms are dependent on the presence of specific viral products, RNA or proteins.

The disruption of PBs during HCV infection relies on an interaction between viral

proteins and PBs components (Ariumi et al., 2011), as well as an alteration of PBs

composition (Perez-Vilaro et al., 2014). HCV hijacks PBs component by redistributing

DDX6, Lsm1, Xrn1, PATL1, and Ago2 to the HCV replication factory localized around lipid

droplets, resulting in enhanced HCV replication. The flavivirus WNV can also co-opt

essential PB proteins, such as Ago2, DDX3, GW182, Xrn1 and Lsm1 for their replication by

re-localising these proteins to a viral replication centre (Chahar et al., 2013; Emara &

Brinton, 2007). An interaction between the viral proteins NS1 with the PBs component

RAP55 also results in inhibition of PB formation during influenza A virus infection (Mok et

al., 2012). Thus, to investigate the mechanisms by which FCV and MNV-1 infection disrupt

PBs, further studies should aim at dissecting the potential re-localization or association of

PBs components within caliciviruses replication complex using immunofluorescence

microscopy studies.

219

It has been shown that the picornavirus 3CPro can disrupt PB formation through the

cleavage of specific PB-nucleating proteins, such as Xrn1, Pan3, and Dcp1a (Dougherty et

al., 2011). We previously suggested that the inhibition of stress granules by FCV infection

might be due to the cleavage of the SG-nucleating protein, G3BP1 by the calicivirus 3CLpro

which has similar targets as the picornavirus 3Cpro (Sosnovtseva et al., 1999). We could

speculate that the repression of PB assembly by FCV and MNV-1 might result from the

cleavage of critical PB components, such as Dcp1. Further studies are therefore required to

investigate the cleavage of Dcp1 or other PBs components using immunoblotting of cell

lysates from FCV and MNV-1 infected cells. In addition, the overexpression of a wild type

FCV or MNV 3CLpro or an inactive mutant, and examination of the effect on PBs, should

indicate whether the calicivirus 3CLpro play a role in PB assembly.

In addition to these mechanisms, signalling pathways have also been linked with the

disruption of PBs. For example, the formation of PBs in response to glucose deprivation in

S.cerevisiae is dependent on the inactivation of the PKA signalling pathway (Ramachandran

et al., 2011). The activation of PKA signalling leads to the phosphorylation of Pat1, a critical

protein for PB assembly, impairing the interaction between Pat1 with other PB components,

such as the RNA helicase Dhh1, and inhibiting the PB assembly (Ramachandran et al., 2011).

As similar behaviour was observed under different stress condition, such as hyperosmotic

stress (Shah et al., 2013). By contrast, the activation of PKA has no effect on stress granule

formation (Shah et al., 2013). Previously, it has been reported that the activation of PKA is

necessary for porcine enteric calicivirus (PEC) replication (Chang et al., 2002). While other

PEC do not replicate in cell culture, the Cowden strain of PEC can replicate in a continuous

cell line (LLC-PK)-containing intestinal content (IC) from uninfected gnotobiotic pigs and

viral replication activates the PKA signalling pathway to inhibit IFN-mediated STAT1

activation (Chang et al., 2002; Chang et al., 2004). Therefore we could speculate that other

220

caliciviruses may also result in activation of PKA, in turns leading to Pat1 phosphorylation

and PBs disruption. To test this hypothesis, further work is required to investigate the

activation of PKA signalling and Pat1 phosphorylation during FCV and MNV-1 infection.

It has been demonstrated that the dissociation of Dcp1a from PBs is strongly

associated with activation of the JNK signalling pathway, part of the MAPK signalling

pathway, leading to the phosphorylation of Dcp1a at the residue S315 (Rzeczkowski et al.,

2011). Moreover, the localisation of DCP1a, Edc4, and Xrn1 to PBs is also affected by JNK

activation in human cells (Rzeczkowski et al., 2011). Previous work from our laboratory has

demonstrated that calicivirus replication is dependent on the activation of two of the main

arms of the MAPK signalling pathway, ERK and p38 (Royall et al., 2015). This activation is

critical to induce eIF4E phosphorylation and mediate host translational control during MNV-

1 infection (Royall et al., 2015). Thus, we could investigate whether calicivirus infection also

activate the remaining arm of the MAPK signalling pathway, namely JNK, using inhibitors of

JNK and immunoblotting studies.

5.5.2 Correlation between PBs and SGs assembly

The results from chapter 3 suggested that FCV infection impairs SGs assembly, while

this chapter revealed that FCV infection also impairs PBs formation. Given that some, but not

all, previous studies proposed that SGs assembly is dependent on pre-existing PBs (Buchan et

al., 2008), we could propose that the inhibition of SGs formation is a consequence of PBs

disruption. However the directional link between PBs and SGs is still controversial. Thus to

investigate the link between PBs and SGs in cells supporting FCV replication we studied the

impact of stress inducing SGs on PBs assembly. The results indicate that hydrogen peroxide,

221

which is potent inducer of SGs (Figure 3.21 & 3.22), disrupt PBs (Figure 5.12). Similarly,

sodium arsenite, a potent inducer of SGs (Figure 3.17 & 3.18), also disrupts PBs although to

a lesser extent (Figure 5.12). Therefore, these results would not support a model in which

SGs assembly is dependent on pre-existing PBs. In contrast, this would support the more

recent model in which mRNAs are found in dynamic equilibrium between polysomes and

cytoplasmic RNA granules (Anderson & Kedersha, 2009b; Kedersha & Anderson, 2009;

Kedersha et al., 2005). By inducing SGs formation with sodium arsenite or hydrogen

peroxide, this would displace the equilibrium in favour of SGs and have a negative knock-on

effect on PBs. However this results are only preliminary an further studies would be needed

to confirm this model, for instance by using stress that promote PBs and analysing the effect

on SGs assembly, or by monitoring the transition of marker mRNAs between the difference

RNA granules in response to stress.

Correlating with the potent effect of hydrogen peroxide on PBs disruption, several

studies have demonstrated an association between hydrogen peroxide treatment and

depolymerisation of microtubules, which is required to deliver mRNAs to PBs (Hu & Lu,

2014; Lindsay & McCaffrey, 2011). In addition, it was proposed that hydrogen peroxide also

plays a role in the phosphorylation of critical PB proteins components (Yoon et al., 2010).

Dcp2 protein is phosphorylated in response to hydrogen peroxide in Saccharomyces

cerevisiae, affect its association with other PB components and impairing their assembly

(Yoon et al., 2010). Furthermore, the hyper-phosphorylation of Dcp1 during cell mitosis also

results in PB inhibition (Aizer et al., 2013).

In conclusion the results from this chapter indicate that caliciviruses infection disrupt

PBs, and suggest several putative mechanisms responsible for this: via the cleavage of the

222

critical PB-nucleation protein, such as Dcp1, via an interaction between viral proteins and PB

components, through phosphorylation certain PB proteins, such as Dcp1 and Pat1 or via the

regulation of a specific signalling pathway, such as PKA or JNK pathways. The inhibition of

PB assembly upon oxidative stress that induce SG formation in the same cell line also lead to

suggest that these granules are formed independently from each other at least in our system.

223

Chapter 6

Conclusion

224

HuNoV is one of the most important causes of acute gastroenteritis worldwide and

responsible for approximately 50% of all gastrointestinal outbreaks, leading to more than

200,000 deaths annually. In the UK, more than a million people are infected every year and

the consequences of this illness amount to an economical loss of more than £100 million per

annum. Viruses of the Norovirus genus are members of the family Caliciviridae, subdivided

into six genogroups (GI-VI) containing several genotypes that infect human and animal hosts.

Due to the lack of suitable cell culture systems to study HuNoV, two animal caliciviruses,

FCV and MNV provide surrogate models to study norovirus biology and interactions with the

infected hosts. Using these models, previous studies have clearly highlighted that calicivirus

have evolved a complex relationship with their hosts to hijack to cellular translation

machinery and mediate viral translation.

In response to stresses such as viral infection, host cells have evolved several

responses leading to the temporary inhibition of protein synthesis to halt viral replication.

One of these responses is mediated by the formation of cytoplasmic RNA granules, SGs and

PBs, where mRNA are either translationally stalled or degraded, respectively. Moreover these

granules are thought to contribute to the immune response against viruses. Therefore, in order

to replicate, viruses have evolved different mechanisms to interfere the assembly of these

cytoplasmic granules. Thus, to better understand the relationship between caliciviruses and

the infected host, I have investigated the formation of RNA granules during caliciviruses

infection using FCV and MNV-1 as a model for HuNoV.

The results discussed in Chapter 3 suggest that infection with either MNV-1 or FCV

did not induce the formation of SGs. These observations however differ between MNV-1 and

FCV. Indeed, I could not detect the formation of SGs in RAW264.7 and BV-2 cells infected

225

with MNV-1 or mock cells that exposed to cellular stress such as sodium arsenite. The

absence of SGs in RAW264.7 and BV-2 cells may be attributed to the inability of these cells

to form stress granules. I propose that this effect may be due to the over-expression of the

protein MCPIP1 protein, which has previously been shown to impair SG formation in

RAW264.7 cells in response to stress or infection. Unlike RAW264.7 and BV-2 cells, CRFK

cells, that support FCV infection, have the ability to form SGs in response to cellular stress.

Furthermore, FCV infection prevents the assembly of SGs in CRFK cells, and actively

disrupts the assembly of SGs formed in response to cellular stresses such as sodium arsenite

and hydrogen peroxide. Moreover, these results suggest that this inhibition of SGs assembly

is associated with viral replication as UV-inactivated virus does not impact on SGs formation.

This implies the involvement of a yet unidentified viral product, RNA or protein, in the

inhibition of SGs response. Based on previous studies, I propose that the viral protease may

contribute to this by cleaving an important SG-nucleating protein or that non-structural

protein may sequester SG components. Further studies will aim at dissecting the molecular

basis of these observations.

Although FCV infection prevents SGs assembly in infected cells, some neighbouring

cells in the vicinity of infected cells displayed SGs, suggesting a cell-to-cell induction

mechanism via a biological or chemical signal explored in Chapter 4. Using virus-free

supernatant from FCV-infected cells, I demonstrated the paracrine induction of SGs

assembly. This is an unprecedented observation that could reflect a new antiviral stress

response. Our results show that this paracrine induction is not mediated by interferon or

soluble RNA molecule. In contrast, heat shock inactivated the supernatant ability to induce

SGs, suggesting that the signalling messenger could have a proteinaceous or more complex

origin.

226

Several studies have suggested that stress granules and perhaps P-bodies, may serve

as sentinel mechanisms for IFN activation by acting as a cytoplasmic platform that enhances

IFN and cytokine signalling through the condensation of IFN signalling moieties with their

ligands within RNA granules. Stress granules induced in a paracrine manner in uninfected

cells may therefore be important for priming neighbouring cells against infection.

Moreover this raises the question whether these granules may help us to develop

novel therapeutic or preventive strategy for viral infectious diseases. As this induction of SGs

prevent viral replication and triggers an antiviral response, the identification of the paracrine

SG inducer could lead to new strategies to block viral replication. As the mechanism targeted

is based on the aggregation of host proteins, this would be unlikely to lead to the

development of drug-resistant viruses. It also raises the possibility of developing novel and

broad antivirals therapeutics with broad spectrum activity.

Finally, in addition to the effect of infection on SGs, we have also analysed the

assembly of PBs during FCV and MNV-1 infection. The results discussed in Chapter 5

showed a strong disruption of PBs assembly during infection by either FCV or MNV-1.

Again, the association of the disruption of these granules with viral replication may also

reflect an important role for viral product in this disruption. This may occur via the cleavage

an essential PB-component by viral protease, an interaction of specific viral protein with

certain PB-component or phosphorylation of critical PB-components by activation a specific

signalling pathways could mediated this mechanisms of disruption. Further work will be

required to dissect the mechanisms leading to the PBs disruption during calicivirus infection.

227

In conclusion, Figure 6.1 summarizes the current working model of caliciviruses

interaction with the mRNA metabolism pathways and RNA granules. In this model, FCV

and MNV-1 infection prevent the inhibition of cellular translation in infected cells by

disrupting the assembly of SGs (FCV) or/and PBs (FCV and MNV). This may prevent the

induction of antiviral responses and maintain viral translation. On the other hand, the

paracrine induction of SGs in bystander cells may reflect a new antiviral pathway induced by

caliciviruses to control infection.

228

Figure 6.1 Proposed working model for the effect of FCV and MNV-1 on stress granules

and P-bodies. Left side shows that the infection prevent the inhibition of cellular translation

via disruption SGs (FCV) or PBs (FCV and MNV-1). Right side shows the paracrine

induction of SGs in bystander cells responding to FCV infection through an unknown

signalling molecule (?).

229

Chapter 7

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Appendix 1

0.1% Triton X-100

- 100 µl Triton X-100

- 100 ml MilliQ water

PBS/BSA

- 1g BSA (Sigma)

- 200 ml autoclaved PBS

4% Paraformaldehyde

- 8g Paraformaldehyde

- 200 ml autoclaved PBS

- 20 µl CaCl2

- 20 µl MgCl2

-

ToPro-3

- 1µl ToPro-3

- 10 ml PBS