MOLECULAR CHARACTERIZATION OF THE NUCLEOCAPSID PROTEIN OF

ARTERIVIRUSES

A Thesis

Presented to

The Faculty of Graduate Studies

of

The University of Guelph

by

HAKIMEH MOHAMMADI

In partial fulfilment of requirements

for the degree

Doctor of Philosophy

January 2010

© Hakimeh Mohammadi, 2010

ABSTRACT

MOLECULAR CHARACTERIZATION OF THE NUCLEOCAPSID PROTEIN OF

ARTERIVIRUSES

Hakimeh Mohammadi Advisor:

University of Guelph, 2010 Dr. Dongwan Yoo

Co-advisor:

Dr. Shayan Sharif

Nuclear localization of the nucleocapsid (N) protein of arteriviruses and the

possible function of N in the nucleus for host cell function modulation were studied.

Subcellular localization of N of lactate dehydrogenase-elevating virus (LDV) was

determined by tagging N with enhanced green fluorescence protein (EGFP) on the N- and

C-termini. Both N-EGFP and EGFP-N fusion proteins were found to localize to the

nucleus and nucleolus of gene-transfected cells. A ‘pat4’ motif was identified in N as a

potential nuclear localization signal (NLS), and its functional significance was

determined by expressing a series of deletion constructs. The results showed that the

‘pat4’ NLS in LDV N was essential for nuclear translocation. LDV N interacted with

importin-α and -β proteins suggesting that its nuclear localization is mediated through the

importin-dependent nuclear transport pathway. This study was expanded to the equine

arteritis virus (EAV) N protein. The EAV N gene was fused with EGFP at the N- or C-

terminus, and its cellular distribution was investigated in gene-transfected cells. Both N-

EGFP and EGFP-N fusion proteins of EAV N accumulated in the nucleus and nucleolus

in addition to the cytoplasm. A series of deletion mutants were made by progressively

deleting amino acids from the C- and N-termini of EAV N to determine the functional

significance of the NLS-motif at amino acids positions 4-16. Studies using the mutant

genes showed that the EAV N nuclear localization required the entire amino acid

sequence from 4-20 position. EAV N interacted with both importin-α and -β proteins but

also with some other fusion proteins suggesting that factors other than the importin-

regulated nuclear transport pathway may be involved in this process. The simian

hemorrhagic fever virus (SHFV) N gene was cloned as a fusion protein with EGFP. The

functionality of the potential ‘pat7’ type NLS present at amino acid positions 22-28 of

SHFV N was determined by constructing a series of deletion mutants and individually

expressing them in cells. The results showed that the ‘pat7’ NLS was essential for nuclear

translocation of SHFV N. The interaction of SHFV N with importin-α suggests that N

recruits this chaperone protein for its nuclear localization. These finding confirm that the

nuclear localization of N is a common property of arteriviruses. Next, modulation of cell

cycle by arteriviruses was investigated. Replication of PRRSV and EAV in cells lead to

accumulation of cells at the G2/M phase. Furthermore, cells expressing N protein from

all four arteriviruses indicated that the number of cells in the G2/M phase was higher

compared to controls. The modulatory effect of N protein on G2/M arrest was similar for

all four N proteins, although PRRSV N and EAV N showed stronger effects on the cell

number increase at the G2/M phase than LDV N and SHFV N. The results obtained from

this research improve our understanding of N protein nuclear localization and its

biological role during infection by members of the Arteriviridae family.

i

ACKNOWLEDGEMENTS

This is an honor for me to thank those who made this thesis possible:

I would like to express my sincere gratitude to my advisor, Dr. Dongwan Yoo, for

providing me with the great opportunity to prepare my research under his supervision in

his laboratory. He has been of such a great help and support throughout in many aspects

including but not restricted to writing grant proposals, scholarship applications and this

thesis. I am grateful to his support, patience and guidance and will always be.

I would like to render my heartfelt appreciations to Dr. Shayan Sharif who

thoughtfully and kindly served as my co-advisor after Dr. Yoo left, providing unfaltering

support and learning opportunity in his lab. From the day one I started my PhD, I have

always relied on his sage hints and discernment. His enthusiasm, inspiration, and great

efforts in explaining complicated matters so perceivably have secured him one of my best

teachers and mentors ever. Throughout the ups and downs of my research and thesis-

writing period as well as the hard times in my life he, with his heart of gold has always

been one of a kind of encouragement, support, initiation, and morale. I would have been

lost without his help in those dire days of dismal doldrums. Moreover by allowing me to

be a part of his laboratory I was privileged to get to know with many nice and talented

people in the laboratory. I doubt that I could ever be able to fully convey my gratitude to

him the way I feel it and hence suffice to round up with saying that I am eternally grateful

to him.

I would also like to thank the other members of my committee, Dr. Dorothee

Bienzle, Dr. Ray Lu, and Dr. Sarah Wootton for the great succor they all provided me

with during this research project. Special thanks go to Dr. Bienzle’s extremely helpful

comments throughout the work, using DAPI in particular as well as creating learning and

training opportunities of professional software and instruments. I am very grateful to Dr.

Wootton who generously not only allowed me to complete my research project after

taking Dr. Yoo’s laboratory over but has also made an invaluable companion, both

professional and confidant.

I wish to thank the members of the Department of Pathobiology including

professors and staff who have helped me from the day one I started my PhD. I would

especially like to thank the Chair of the Department Dr. Robert Jacobs for his kindness

and help at the gloomy times of despair. Without his full support and management I

would have not been able to complete my degree. My especial thanks go to Betty-Anne

McBey who has been always helpful in FACS area and without her help and effort

nothing can be done there. Donna Kangas, Jean Bagg, Cathy Bernardi, Elizabeth

Gilbertson, and many others deserve special mention.

I would also like to thank my old colleagues in Dr. Yoo’s past laboratory

including Frances Lai, Paul Rosenfeld, Dr. Yanlong Pei, Cheng Song, Dr. Gang Li, Dr.

ii

Changhee Lee, Sheila Watson, and Maged Gomaa for their friendship, help and

willingness to participate in the scientific environment we had in the laboratory.

I would like to thank to all friends and colleagues in Dr. Sharif’s lab who so

warmly welcomed, supported and treated me that the lab time became an enjoyable span

of day: Leah Read, Hamid Haghighi, Payvand Parvizi, Niroshan Thanthrige, Kamran-Ul

Haq, Raveendra Kulkarni, Jen Brisbin, Amirul Mallick, and other members of this lab.

During my PhD at Guelph, I have met many wonderful people in the Department,

OVC, and university of Guelph who have offered their unrestrained friendship and help. I

am especially thankful for the International Advisor Mr. Benny Quey who has been a full

support whenever needed. I am so blessed to have so many friends and family members

who have been all help and for them I am always grateful.

I would also like to thank my and my husband’s families for the support they have

provided me through my entire life. Particularly I would like to thank my parents

Mohammadali Mohammadi and Sara Hosaini for their unconditional love and support

from the day I was born. Without their support and love I would not be able to stand at

the place I am in now. I would also like to thank my brothers and sisters, and my brother-

and sister- in law. They have always been inspiring me with love and support. I would

also very much like to remember the dear memories and supports of my departed brother

and mother-in-law who always loved and encouraged me to study and overcome the

difficulty that life might throw in.

And the last but most importantly, my great appreciation is for the greatest ocean

of affection, love and support; my loving husband, Alireza Omumi, who has been my

best friend and comrade in bad and good times. Without your passionate love, support,

encouragement and inspiring assistance, I would not be able to complete this thesis. I am

here now because you have always been inspiring, supportive and patient. I am forever

grateful to your kind heart.

In conclusion, I recognize that this research would not have been possible without

the financial assistance of “Iranian Ministry of Health and Medical Education” that

provided me the full scholarship for 4 years and Dr. Yoo’s funds (Natural Sciences and

Engineering Research Council, and US Department of Agriculture National Research

initiative), as well as the support of the department of pathobiology and university of

Guelph. I express my gratitude to those.

iii

I wish to dedicate this thesis to my departed brother, Abdolkarim Mohammadi.

iv

TABLE OF CONTENTS

ACKNOWLEDGEMENTS...…..…………………………………………………………i

TABLE OF CONTENTS……………………………………….………………………..iii

LIST OF TABLES…………………………………………………….………………….vi

LIST OF FIGURES……………………………………………………………………...vii

LIST OF ABBREVIATIONS……………………………………………………..…...…ix

INTRODUCTION……………………………...…………………………………………1

CHAPTER 1:

I. Review of Literature

1. The family Arteriviridae, Classification, and the Diseases………….…2

1.1 History and Taxonomy

1.2 Diseases Caused by Arteriviruses

1.3 Transmission

2. Virion Structure and Genome Organization of Arteriviruses…………..4

2.1 Virus Particle and Virion Composition

2.2 Genome Organization

2.3 Major Structural Proteins

2.3.1 M protein

2.3.2 Glycoprotein 5

2.3.3. Nucleocapsid (N) protein

2.4 Minor Structural Proteins

3. Life Cycle of Arteriviridae……………………………………………12

3.1 Cell Tropism

3.2 Attachment and Entry

3.3 Replication and Transcription

3.4 Virion Assembly and Release from Cell

4. Pathogenesis of Arteriviruses…………………………………………21

4.1 Natural Infection and Persistence

4.1.1 Equine Arteritis Virus (EAV)

4.1.2 Porcine Reproductive and Respiratory Syndrome

Virus (PRRSV)

4.1.3 Lactate Dehydrogenase Elevating Virus (LDV)

4.1.4 Simian Hemorrhagic Fever Virus (SHFV)

4.2 Immune Response to Arterivirus Infections.

4.2.1 Innate Immune Response to Arteriviruses

4.2.2 Adaptive Immune Response to Arteriviruses

5. Nuclear Localization of Viral Proteins in +ssRNA viruses……..…….28

v

5.1 Nucleolus and Nuclear Localization of Proteins

5.2 Nucleolar Localization of +ssRNA Virus Proteins

5.3 Function of Nuclear Localization of Viral Nucleocapsid

Proteins

5.4 Modulation of Cell Cycle by Viral Proteins

5.5 Nucleocapsid Protein in Arteriviridae

II. Objectives of the Thesis………………………………………………………44

CHAPTER 2: The Lactate Dehydrogenase-Elevating Virus Capsid Protein is a Nuclear-

Cytoplasmic Protein.

Abstract…………………………………………………………………………..47

Introduction………………………………………………………………………48

Materials and Methods…………………………………………………………...50

Results……………………………………………………………………………54

Discussion………………………………………………………………………..64

CHAPTER 3: Nuclear Localization of Equine Arteritis Virus and Simian Hemorrhagic

Fever Virus Nucleocapsid Proteins.

Abstract…………………………………………………………………………..69

Introduction………………………………………………………………………70

Materials and Methods………………………………………………………...…72

Results……………………………………………………………………………80

Discussion………………………………………………………………………..96

CHAPTER 4: The Role of Nucleocapsid Protein of Arteriviruses in Cell Cycle

Modulation.

Abstract…………………………………………………………………………102

Introduction……………………………………………………………………..103

Materials and Methods………………………………………………………….105

Results…………………………………………………………………………..108

Discussion………………………………………………………………………114

CHAPTER 5: General Conclusions……………………………...……………………..119

REFERENCES…………………………………………………………………………124

vi

LIST OF TABLES

CHAPTER 2: The lactate dehydrogenase-elevating virus capsid protein is a nuclear-

cytoplasmic protein.

Table 2.1. List of primers used for EGFP fusion constructions for LDV N……………..51

Table 2.2. Site-directed mutagenesis of LDV N NLS motif and subcellular localization of

NLS mutants……………………………………………………………………………..60

CHAPTER 3: Nuclear localization of equine arteritis virus and simian hemorrhagic

fever virus nucleocapsid proteins.

Table 3.1. List of primers used for EGFP fusion constructions for EAV N…………….74

Table 3.2. List of primers used for EGFP fusion constructions for SHFV N…………...75

Table 3.3. Site-directed mutagenesis of the SHFV N NLS motif and subcellular

localization of NLS mutants……………………………………………………………..93

vii

LIST OF FIGURES

CHAPTER 1: Literature review

Figure 1.1: Schematic representation and genome organization of arteriviruses…………5

Figure 1.2: Overview of the genomes of the arteriviruses………………………………...7

Figure 1.3: Schematic representation of the life cycle of arteriviruses…………………..16

Figure 1.4: The nested set of RNAs in arteriviruses and the discontinuous strategy for

mRNA transcription………...…..…………….………………………………………….19

Figure 1.5: The schematic illustration of the nuclear protein transport …………………30

Figure 1.6: Schematic presentation of a cell cycle……………………………...……….38

CHAPTER 2: The lactate dehydrogenase-elevating virus capsid protein is a nuclear-

cytoplasmic protein.

Figure 2.1: Subcellular localization of LDV-N protein………………………………….55

Figure 2.2: Identification of an NLS motif in the LDV-N protein and its functional

mapping…………………………………………………………………………………..58

Figure 2.3: Subcellular localization of NLS mutants…………………………………….61

Figure 2.4: Subcellular localization of the NLS-null LDV-N-EGFP mutants in HeLa and

NIH-3T3 murine cells…………………………………………………………………....62

Figure 2.5: Interactions of LDV-N with mouse importin proteins by GST-pull down

assays…………………………………………………………………………………….65

CHAPTER 3: Nuclear localization of equine arteritis virus and simian hemorrhagic

fever virus nucleocapsid proteins.

Figure 3.1: Confirmation of the EAV N nuclear localization ……………………….......81

Figure 3.2: Subcellular localization of EAV N protein……………………………….....82

Figure 3.3: Identification of NLS in EAV N protein and its functional mapping……….84

Figure 3.4: Subcellular localization of EAV N NLS mutants…………………………...86

Figure 3.5: Interaction of EAV-N with importin proteins by GST-pull down assay……87

viii

Figure 3.6: Subcellular localization of SHFV-N protein………………………………...89

Figure 3.7: Identification of NLS in SHFV N protein and its functional mapping……...90

Figure 3.8: Subcellular location of the NLS-null SHFVN-EGFP mutant……………….94

Figure 3.9: Interaction of SHFV N with importin proteins by GST-pull down assay…...95

CHAPTER 4: The role of nucleocapsid protein of arteriviruses in cell cycle modulation.

Figure 4.1: Experimental design for analysis of Arteriviruses and their nucleocapsid

protein effects on cell cycle…………………………………………………………….107

Figure 4.2: Co-localization of LDV N and EAV N with fibrillarin…………………….109

Figure 4.3: CPE of PRRSV infected MARC-145 cell at different times ………………111

Figure 4.4: Cell cycle profile of Mock/PRRSV infected MARC cells…………………112

Figure 4.5: Cell cycle profile of mock/EAV infected BHK-21 cells…………………...113

Figure 4.6: Cell cycle profile in arterivirus N transfected BHK-21cells…….………....115

ix

LIST OF ABBREVIATIONS

BHK Baby hamster kidney

BSA Bovine serum albumin

CAS Cellular Apoptosis Susceptibility protein

Ci Curie

CPE Cytopathogenic effects

DAPI 4',6-Diamidino-2-phenylindole dihydrochloride

DFC Dense fibrillar component

DMEM Dulbecco’s minimum essential medium

DNA Deoxyribonucleic acid

dpi Day post infection

EAV Equine arteritis virus

E.coli Escherichia coli

EDTA Ethylenediamine tetraacetic acid

EM Electron microscopy

ER Endoplasmic reticulum

FBS Fetal bovine serum

FACS Fluorescence activated cell scanning

FC Fibrillar center

G1 First gap

G2 Second gap

GC Granular component

GDP Guanosine diphosphate

GP Glycoprotein

RanGEF Ran Guanine nucleotide -Exchange Factor

GST Glutathione-S transferase

GTP Guanosine-5'-triphosphate

h Hour

HCV Hepatitis C virus

HDV Hepatitis delta virus

hpi Hours post infection

HIV Human immunodeficiency virus

IBV Infectious bronchitis virus

ID50 Infectious dose 50%

IF Immunofluorescence

IgG Immunoglobulin G

Impα Importin α

Impβ Importin β

IPTG Isopropyle-β-D-thiogalactopyranoside

IRES Internal ribosome entry site

kDa Kilodalton

JEV Japanese encephalitis virus

JSs Junction sites

LD50 Lethal dose 50%

LDV Lactate dehydrogenase-elevating virus

x

LV Lelystad virus

M Mitosis

M protein Matrix protein

μl Microliter

ml Mililiter

MAb Monoclonal antibody

MHV Mouse hepatitis virus

MOI Multiplicity of infection

mRNA Messenger RNA

N Nucleocapsid protein

NE Nuclear envelope

NES Nuclear export signal

NLS Nuclear localization signal

NoLS Nucleolar localization signal

NPC Nuclear pore complex

Nsp Nonstructural protein

ORF Open reading frame

PAGE Polyacrylamide gel electrophoresis

PAMS Porcine alveolar macrophages

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PFU Plaque forming unit

PMSF Phenylmethylsulfonyl fluoride

PRRSV Porcine reproductive and respiratory syndrome virus

Ran Ras-related nuclear protein

RanGAP Ran GTPase activating protein

RIs Replicative intermediates

RNA Ribonucleic acid

rRNA Ribosomal RNA

RT Reverse transcriptase

SARS Severe acute respiratory syndrome

SDS Sodium dodecyl sulfate

sgmRNA Subgenomic mRNA

SHFV Simian hemorrhagic fever virus

ssRNA Single stranded RNA

tRNA Transfer RNA

TCID50 50% tissue culture infectious dose

TGEV Transmissible gastroenteritis virus

UBF upstream binding factor

TLRs Transcription regulating sequences

UTR Untranslated region

1

INTRODUCTION

Arteriviridae is a family of viruses comprising four viruses some of which cause

important diseases in animals with major economic losses. These viruses are: equine

arteritis virus (EAV), lactate dehydrogenase-elevating virus (LDV), porcine reproductive

and respiratory syndrome virus (PRRSV), and simian hemorrhagic fever virus (SHFV).

These viruses have a linear positive stranded RNA as genome and based on similarity of

their genome organization and replication strategy with Coronaviridae are categorized in

Nidovirales order of viruses. They replicate in the cytoplasm of infected cells but it has

been reported that the nucleocapsid protein of PRRSV and EAV, as one of the major

structural proteins can localize in the nucleus of infected cells and interact with nucleolar

proteins involved in cellular activities. A similar pattern of cellular distribution has been

reported for the nucleocapsid protein of coronaviruses. Understanding the nucleolar

localization of N protein of other members of arteriviruses demands further investigation

into its biological function and whether this is a common feature of members of this

family in the Nidovirales order. Moreover, previous studies of coronavirus N proteins and

their modulation of cell cycle have raised the possibility that the capsid protein of

Arteriviridae has a major role in cell cycle progression during virus replication. Thus,

elucidation of nuclear localization of capsid proteins of all four member viruses of the

Arteriviridae family and involvement in the cell cycle could open a window to

understand the importance of this feature in virus pathogenesis. Moreover, advancing our

understanding of viral N protein interactions with host proteins may lead to the design of

novel vaccines. The new strategies for vaccine against arteriviruses could target the N

protein as one of the most important viral proteins to achieve better prevention.

2

CHAPTER 1

I. REVIEW OF LITERATURE

1. Arteriviridae, Classification, and the Diseases

1.1 History and Taxonomy. The family Arteriviridae was established by the

International Committee on Txonomy of Viruses (ICTV) in 1996 (Cavanagh et al., 1994;

Cavanagh 1997). Arteriviridae is comprised of four member viruses: equine arteritis virus

(EAV), lactate dehydrogenase-elevating virus (LDV), porcine reproductive and

respiratory syndrome virus (PRRSV), and simian hemorrhagic fever virus (SHFV). Three

of these viruses, EAV, LDV, and SHFV have been known for the last 4-5 decades,

whereas PRRSV emerged in the late 1980s in Europe and early 1990s in North America

(Snijder and Meulenberg, 1998; Albina, 1997). These viruses cause different diseases in

susceptible animals, ranging from mild asymptomatic to persistent or severe clinical

syndromes. EAV is the prototype virus of the family and was first isolated in 1957 in

Bucyrus, Ohio (USA) from the lungs of horses suffering from abortion (Doll et al, 1957).

PRRSV is the causative agent of porcine reproductive and respiratory syndrome (PRRS)

(Collins et al., 1992; Rossow, 1998). LDV is the causative agent of an asymptomatic

persistent infection with lifelong viremia in mice. LDV has been mostly used as an in

vivo model in many investigations on persistent infections (Riley et al, 1960; Rowson and

Mahy, 1985). SHFV is the etiological agent of a fatal hemorrhagic fever of monkeys and

was first isolated in 1968 during several outbreaks of a febrile hemorrhagic disease in

colonies of macaque monkeys at the National Institute of Health (NIH) (Palmer at al,

1968; Allen et al, 1968; Tauraso et. al 1968). Although SHFV leads to 100% mortality in

3

macaque, it has been shown to cause an endemic asymptomatic infection in some other

genera of African monkeys (London et al, 1977; Gravell et al, 1986b).

A unique feature of the family Arteriviridae is the production of a nested set of

subgenomic (sg) mRNAs from their genome, a property shared with another family of

viruses, the Coronaviridae. Based on the similarity of their genome structure, mechanism

of gene expression, and replicase genes, Arteriviridae and Coronaviridae have been

placed in the new order of viruses, Nidovirales (the nidus in the Latin means the “nest’),

along with Roniviridae, a newly identified group of viruses in invertebrates (de Vries et

al., 1997).

1.2 Diseases caused by arteriviruses. The disease caused by EAV may be asymptomatic

or symptomatic with severe manifestations including edema, hemorrhage of small

arteries (hence the name EAV), and abortion in pregnant mares (Doll et al, 1957, and Del

Piero, 2000). PRRS is one of the most economically important viral diseases of the swine

industry. Manifestations of PRRSV-induced disease include severe reproductive failure

in sows and gilts with a high abortion rate and an increased number of stillborn and weak

piglets. The disease has also been related to respiratory problems due to interstitial

pneumonitis (Rossow, 1998). LDV infection in mice has no apparent clinical signs except

an increased activity of lactate dehydrogenase in plasma, from which the name of LDV is

derived (Plagemann et al., 1995). LDV has very limited in vitro replication with restricted

tropism to mouse primary macrophages (Brinton-Darnell et al., 1975; Brinton-Darnell

and Plagemann 1975; Stueckemann et al., 1982; Onyekaba et al., 1989a). LDV infection

is usually asymptomatic, but in specific strains of mice it has been shown to cause lethal

poliomyelitis due to cytocidal replication of the virus (Contag and Plagemann, 1989).

4

SHFV induces a lethal febrile hemorrhagic disease in different genera of Asian and

African monkeys (Palmer et al., 1968; Tauraso, et al., 1968).

1.3 Transmission. Both EAV and PRRSV can be spread mainly through the respiratory

system and both viruses can induce persistent infections in animals. Persistent infection

in male animals enables venereal transmission of the virus a common way for spread of

EAV and PRRSV (Timoney and McCollum 1993; Wensvoort et al., 1993). PRRSV has

been found in saliva, respiratory tract secretions, and urine. Also, the virus can be

transmitted via the placenta (Rossow, 1998). LDV is secreted in urine, feces, and saliva,

and horizontal transmission is not an effective route of LDV spread except in invasive

fighting in male mice. The transmission of LDV through placenta and milk is a more

effective route of virus transfer (Plagemann and Moennig, 1992). SHFV can transmit

through direct contact and aerosols, but unlike other members of the family, it is not

transmitted vertically via placenta to the fetus (Gravell et al., 1986a).

2. Virion Structure and Genome Organization of Arteriviruses

2.1 Virus Particle and Virion composition. Arteriviruses are a group of spherical

viruses with virion size of 45-60 nm in diameter (Fig. 1.1 a; Brinton, 1999; Snijder and

Meulenberg, 1998; Snijder and Spaan, 2007). Their core is an icosahedral structure of 25-

35 nm enclosed in a lipid envelope with small projections as viral proteins, which are

quite different from the long spikes seen in coronaviruses. These viruses are sensitive to

non-ionic detergents, pH > 7.5 and < 6, and storage at 4oC or higher temperature

(Horzinek et al., 1971; Brinton-Darnell and Plagemann, 1975; Sagripanti, 1984;

Meulenberg et al., 1995a; Burki, 1966; Tauraso et al, 1968; Snijder and Meulenberg,

1998).

5

Fig. 1.1. (A) Schematic representation of PRRSV particle. The virion of arteriviruses is

spherical surrounding an icosahedral capsid formed by the nucleocapsid protein (N)

which in turn encloses the positive sense RNA genome of the virus. The membrane is

composed of five proteins: the membrane protein (M), the major glycoprotein (GP)5, the

minor glycoproteins GP2a and GP4, and the envelope (E) protein. The GP3 is believed to

be structural for the European PRRSV and a secretory protein in case of the North

American type of this virus (Adapted from Dea et al, 2000). (B) The genome

organization of EAV, prototype of the Arteriviridae family of viruses, and transcription

of subgenomic mRNAs (adapted from Snijder and Meulenberg, 1998).

6

Arteriviruses have a buoyant density of 1.13 to 1.17 g/cm3 in sucrose and their

sedimentation coefficient varies from 214 S to 230 S.

A schematic figure of these viruses is shown in Fig. 1.1 a. The nucleocapsid of the

family Arteriviridae contains a positive-sense single-stranded RNA (ssRNA) genome of

12.7 to 15.7 kb enclosed by the nucleocapsid protein (N) (Brinton et al, 1999, Snijder and

Meulenberg, 1998; Snijder and Spaan, 2007). Studies on EAV and PRRSV have

suggested that there are six envelope proteins embedded in the lipid bilayer envelope of

arteriviruses. For all members of the family, the three major structural proteins that have

been described as virion components are the nucleocapsid protein N (12-15 kDa), the

non-glycosylated membrane protein M (16-20 kDa) and the major glycoprotein (GP) (22-

44 kDa, GP5 in PRRSV, LDV and EAV and GP7 in SHFV). M and GP5 of PRRSV, EAV

and LDV form a heterodimer via a disulfide linkage (de Vries et al., 1995a; Faaberg et

al., 1995; Snijder et al., 2003). The major structural proteins are encoded by open reading

frames (ORF) located at the 3’ end of the viral genome (Snijder and Spaan, 2007). The

minor structural proteins include GP2 (coded by ORF2a of PRRSV and ORF2b of

EAV/LDV), GP3, GP4, and the small non-glycosylated envelope protein E (Snijder and

Spaan, 2007). GP2, GP3, GP4 of PRRSV and EAV have been shown to form heterotrimers

in the virus particle (Wieringa et al., 2003; Wissink et al., 2005). Using reverse genetics,

it has been shown that all major and minor structural proteins are necessary in order to

produce infectious EAV (Wieringa et al., 2004).

2.2 Genome Organization. Viruses in the family Arteriviridae contain a single-stranded

positive-sense RNA genome with a length of 12.7 to 15.7 kb (Snijder and Swaan, 2007).

7

Fig. 1.2. Overview of the genomes of the arteriviruses. The genome organization of the

family prototype EAV is shown at the top. The genome includes the replicase open

reading frames (ORF) 1a and 1b which are followed by ORFs 2-7 (9 in SHFV) encoding

for structural proteins. The structural genes include genes for the E protein, three minor

glycoproteins (GP2a/b-4), major glycoprotein GP5 (GP7 in SHFV), and the genes for the

membrane (M) and nucleocapsid (ORF7) proteins. The 3’ end of the SHFV genome has a

large insertion which is assumed for the putative extra ORFs (Snijder and Spaan, 2007).

8

The genome of SHFV has been reported to have a type I cap structure (Sagripanti et al.,

1986). The necessity of cap structure for in vitro transcription of infectious viral RNA

from the full-length cDNA clones of PRRSV and EAV has been described. The full-

length genomes of all members including two European and North American isolates of

EAV, more than 20 European and North American isolates of PRRSV, LDV strains, and

one isolate of SHFV have been sequenced (Fig. 1.2) (Snijder and Spaan, 2007).

Approximately three-fourth of the 5’ end of the genome is comprised of two large ORFs,

called ORF1a and ORF1b, which encode non-structural proteins. The ORF1a-encoded

polyprotein is more variable in sequence and size among members of the family

compared to the ORF1b-encoded polypeptide (Snijder and Meulenberg, 1998). The 3’

part of the genome of EAV, LDV, and PRRSV includes a group of seven smaller

overlapping ORFs which encode structural proteins of these viruses. SHFV genome

contains three additional ORFs at the 3’ end. These ORFs are located between ORF1b

and ORF5 of the other three members of the Arteriviridae family (Fig. 1.2). The

significance of these extra ORFs in the SHFV genome is not clear. In the 3’ end of the

replicase gene and in the leader sequence of EAV, PRRSV, and SHFV, the presence of

small ORFs have been reported with no clear significance (Snijder and Meulenberg,

1998).

2.3 Major Structural Proteins. The three most abundant viral proteins in arteriviruses

are GP5 (GP7 in SHFV), M, and N. These proteins constitute 90-95% of the structural

proteins of the virion and are expressed from three ORFs located at the 3’ end of the

genome in the mentioned order (Snijder and Spaan, 2007).

9

2.3.1 M Protein. The M protein is a non-glycosylated structural protein of arteriviruses

and is known to be one of the most abundant structural proteins. The M protein is the

most conserved structural protein in this group of viruses. Its configuration and function

are similar to the M protein of Coronaviridae and is suggested to be a membrane protein

(de Vries et al., 1992; Kuo et al., 1992; Meulenberg et al., 1993b). The N-terminal half of

the arterivirus M protein is thought to span the membrane three times (de Vries et al,

1992; Faaberg and Plagemann, 1995; Meulenberg et al., 1993b) resulting in a

configuration consisting of the N-terminus of the M protein at the external and the C-

terminus of M towards the internal sides of the virion as well as exposure of a short

stretch of amino acids (10-18 residues) at the surface of the virion (de Vries et al., 1995a;

Faaberg et al., 1995a; Mardassi et al., 1996). It is proposed that the M protein of

Arteriviridae has an important role in assembly and budding of viruses (Wieringa et al.,

2004). The M protein is localized in the endoplasmic reticulum (ER) and through the Cys

residues of its N-terminal ectodomain binds to the Cys residues of GP5, leading to the

formation of heterodimer of M-GP linked by disulfide bonds (de Vries et al., 1995a;

Faaberg et al., 1995a; Mardassi et al., 1996). The disulfide linked M-GP5 heterodimers

have been detected in PRRSV and EAV infected cells (de Vries et al., 1995a). It has also

been reported that disruption of this disulfide bond by treating LDV virion with DTT

decreases the infectivity of this virus. This finding indicates the importance of M-GP5/7

heterodimer in virus infectivity likely at the beginning of the replication cycle (Faaberg et

al., 1995a).

2.3.2 Glycoprotein5/Glycoprotein7 (GP5/GP7). GP5/GP7 is the major GP of

Arteriviridae and is a hydrophobic protein. The internal part of GP5 is presumed to

10

traverse the viral membrane three times (Snijder and Meulenberg, 1998). The size of the

ectodomain is smaller in PRRSV and LDV compared to the EAV equivalent. However, a

common feature in all three viruses is the presence of an N-glycosylation site in

ectodomain. Glycosylation of GP5 of EAV and LDV, is the result of adding different

numbers of repeated lactosamines (de Vries et al., 1992; Li et al., 1998). Neutralizing

antibodies are mainly produced against GP5 in mice and horses infected with LDV and

EAV, respectively (Cafruny et al., 1986; Chirnside et al., 1995a, 1995b). Moreover,

monoclonal antibodies (MAbs) against PRRSV GP5 have been found to neutralize

PRRSV infection (Pirzadeh and Dea, 1997). Reports have shown that the ectodomain of

GP5 is the target for neutralizing antibodies in EAV-infected horses and for MAbs raised

against GP5 (Chirnside et al., 1995a; Balasuriya et al., 1997). The reports also show that

mutations or deletions of this portion of GP5 help EAV mutants escape the host immune

response without affecting their infectivity (Balasuriya et al., 1995; Glaser et al., 1995;

Balasuriya et al., 1997). A similar location of the epitope for neutralizing antibodies has

been shown in the ectodomain of the GP5 glycoprotein of LDV (Li et al., 1998). The

number of glycosylation sites at the ectodomain of GP5 is different in LDV strains

(Faaberg et al., 1995b). This difference is suggested to be involved in the persistence and

neurovirulence of LDV. Li et al. (1998) demonstrated that binding of neutralizing

antibodies to the highly glycosylated GP5 of LDV is decreased.

2.4 Minor Structural Proteins. Several minor structural proteins have been

characterized for arteriviruses. The ORF2a of PRRSV and ORF2b of EAV/LDV encode

GP2 (previously called GPs for EAV). GP2 is a part of the virion in PRRSV and EAV. It

is a type I integral membrane protein and has a signal peptide at the N-terminus and a

11

transmembrane domain located at the C-terminus. This protein is suggested to be N-

glycosylated (de Vries et al., 1992; Meulenberg and Petersen-den Besten, 1996). There

are cysteine residues of EAV GP2, which form not only intrachain disulfide bonds in

GP2-GP2 homodimers found in the virion, but also intermolecular disulfide bonds with

the ectodomain of another minor GP (GP4), resulting in the formation of GP2-GP4

disulfide-linked complexes (Wieringa et al., 2003). The GP2 protein of PRRSV and EAV

remains in the ER when expressed as a recombinant protein (de Vries et al., 1995b;

Meulenberg and Petersen-den Besten, 1996). This suggests that it needs other structural

proteins or assembly of the virion for translocation from the ER to the Golgi complex.

Recently, an ion channel function for the PRRSV E protein has been reported which

suggests that the E protein might have a function in fusion of the virus envelope with the

endosomal membrane leading to disassembly of the genome and nucleocapsid of PRRSV

after entry (Lee and Yoo, 2006). GP3 and GP4 are the glycoproteins encoded by ORF3

and ORF4 of PRRSV, EAV, and LDV, respectively. GP3 is an integral membrane protein

which is highly glycosylated. This glycoprotein is one of the minor structural proteins of

the PRRSV virion (van Nieuwstadt et al., 1996), but it has yet to be detected in EAV and

LDV virions or cells infected with these viruses. The GP3 and GP4 proteins of LDV

have been shown to be soluble and membrane-attached, respectively, in vitro (Faaberg

and Plagemann, 1997). GP4 is also a type I integral membrane protein. MAbs raised

against PRRSV ORF4 protein have been shown to possess neutralizing activity and the

MAb-binding site has been reported to be a highly heterogeneous region of this protein in

different strains of PRRSV (van Nieuwstadt et al., 1996; Meulenberg et al., 1997).

Heterotrimers of GP2, GP3, and GP4 have been reported for EAV (Wieringa et al., 2003).

12

There are three more ORFs located at the 3’ end of SHFV genome which are suggested to

be replicas of ORFs 2 to 4 (Godeny et al., 1998).

3. Life Cycle of Arteriviridae

3.1 Cell Tropism. Viruses in the Arteriviridae family have limited growth ability in

vitro. Primary macrophages from natural hosts are the select cells for their replication.

LDV has a very limited growth in murine macrophage, and it only replicates in specific

primary macrophages of mice (Plagemann and Moening, 1992). SHFV and PRRSV

replicate in simian and porcine macrophages, respectively. They both replicate in African

green monkey kidney cells (MA-104) and its derivatives (CL2621 and MARC-145)

(Gravell et al, 1986b; Benfield et al, 1992; Kim et al, 1993). Some strains of PRRSV

replicate better in porcine alveolar macrophages (PAMs) while some others grow better

in CL2621 (Bautista et al, 1993). Unlike other arteriviruses, EAV infects and replicates in

kidney cell lines such as baby hamster kidney (BHK-21; Hyllseth, 1969), rabbit kidney

(RK-13; Snijder and Meulenberg, 1998), and African green monkey kidney (Vero cells;

Konishi et al, 1975) as well as in primary macrophages. Using one-step growth

experiments, it has been shown that it takes 10 to 20 hrs to detect the first virus particles

released from virus-infected cells (Stueckemann et al., 1982; van Berlo et al., 1982;

Gravell et al., 1986b; Kim et al., 1993). High rates of cytolysis and cell death result from

infection of primary macrophages and cell lines by these viruses. The titer of PRRSV and

SHFV in the cell culture can be 106-10

7 of 50% tissue culture infectious dose per ml

(TCID50 /ml) but it can reach 108

TCID50/ml for EAV (Tauraso et al., 1968; Onyekaba et

al., 1989b; Snijder and Meulenberg, 1998). Replication of LDV in cell culture is very

limited and thus LDV is generally titrated in vivo in mice (Plagemann and Moennig,

13

1992). In cells infected with EAV, SHFV, and PRRSV, cytopathic effects (CPE) are

described as cell rounding and detachment from the surface of culture plates (Hyllseth,

1969; van Berlo et al., 1980; Gravell et al., 1986b; Benfield et al., 1992; Kim et al., 1993;

van Nieuwstadt et al., 1996). Presence of “double membrane vesicles” (DMVs) in

infected cells is common in Arteriviridae, and occurs 3-6 hr post-infection (Snijder and

Meulenberg, 1998; Wood et al., 1970; Stueckemann et al., 1982; Rossow, 1998; Pol et

al., 1997). The role of these structures in replication and viral RNA transcription is not

yet known, though it does not seem that they are involved in assembly of the virus.

3.2 Attachment and Entry. Transfection of non-permissive cells with LDV and PRRSV

genomic RNA leads to production of virus particlest (Inada et al, 1993; Meulenberg et al,

1998). When different cell lines derived from rats and mice were first infected with

murine leukemia virus, they supported later infection with LDV (Inada and Yamazaki,

1991). This suggests that one of the determinants in susceptibility of cell lines to

arteriviruses replication is the existence of specific receptors on the surface of these cells.

The cell tropism has been well investigated for PRRSV. It has been found that pre-

incubation of PRRSV or MARC-145 cells with heparin or cells with heparinase, prevents

PRRSV infection. This suggested that there is a heparin-like molecule on the surface of

MARC-145 cells which supports virus attachment and subsequent entry of the virus (Jusa

et al, 1997). Subsequently, sialoadhesin (or sialic acid-dependent lectin-like receptor 1;

Siglec-1), which is a macrophage-specific molecule, was found to mediate PRRSV

attachment to the surface of alveolar macrophages (Vanderheijden et al, 2003). In other

investigations, heparin sulfate on the surface of macrophages and sialic acid on the

PRRSV virion were shown to play a role in virus entry (Delputte et al., 2002;

14

Vanderheijden et al., 2001; Vanderheijden et al., 2003). In the present model of infection

of macrophages with PRRSV, it is suggested that the virus first binds to heparin sulfate.

This facilitates binding of viral ligands to Siglec-1 which happens via sialic acid present

on viral glycoproteins (Delputte et al, 2005). This is followed by internalization of virus

through endocytosis which is dependent on clathrin-coated vesicles (Fig. 1.3) (Delputte et

al., 2004, 2005, 2007; Delputte and Nauwynck, 2004). As mentioned, infection of the

permissive cells such as MARC-145 is partially mediated by the presence of a heparin-

like molecule on the surface (Jusa et al, 1997). However, the absence of sialoadhesin on

these cells and on cells belonging to non-macrophage lineages, suggested that there was

another molecule which mediated viral entry. It was shown that the micro-filament

vimentin in MARC-145 cells binds to the nucleocapsid protein of PRRSV. This

suggested that vimentin may transport PRRSV to host cells by its interactions with other

compartments of the cytoskeleton (Kim et al, 2006). CD151 has also been shown to

interact with PRRSV RNA and it is suggested to mediate fusion of viral membrane and

endosome (Shanmukhappa et al., 2007). Another investigation showed that scavenger

receptor CD163, a member of the scavenger receptor cysteine-rich family class B, is

necessary for infection of MARC-145 by PRRSV. Its expression in a non-permissive cell

line resulted in replication of PRRSV in these cells (Calvert et al., 2007). A recent study

showed that sialoadhesin is the cellular receptor involved in PRRSV internalization and

CD163 is needed for viral entry (Van Gorp, et al, 2008).

Studies using drugs affecting the pH of intracellular compartments have revealed

that low pH is needed for PRRSV entry. Electron microscopy studies have shown that

PRRSV and LDV particle are visible in clathrin coated vesicles (Kreutz and Ackermann,

15

1996, Kowalchyk and Plagemann, 1985). A recent investigation suggests that EAV also

enters cells by clathrin-dependent endocytosis and is delivered to acidic endosomal

compartments (Nitschke et al, 2008). All the above evidence indicates that arteriviruses

use the low pH-dependant endosomal pathway to enter cells.

3.3 Replication and Transcription. Replication of arteriviruses takes place in the

cytoplasm of infected cells (Fig. 1.3). After virus entry and uncoating, viral genome RNA

is released into the cytoplasm and translated into two large polyproteins, pp1a (1727 for

EAV and 2502 amino acids for PRRSV) and pp1ab (3175 for EAV 3959 amino acids for

PRRSV) which provide the replicase proteins needed for viral RNA synthesis

(Molenkamp et al, 2000b; Snijder and Spaan, 2007). ORF1b of the replicase gene starts

at the -1 position to the stop codon location of ORF1a. The translation of the replicase

polyproteins starts at the 5’ end non-translated region (NTR) of the genome after

“conventional” scanning of the genomic 5’ NTR by ribosome (Van den Born, 2005). The

translation of ORF1b requires two structural signals, located at the ORF1a/1b junction,

which mediate the ribosomal frameshift. One signal is a “slippery” sequence at the

location of the frameshift before the stop codon of ORF1a. The proposed slippery site for

EAV starts upstream of the stop codon of ORF1a (5’GUUAAAC 3’), but for LDV and

PRRSV, there is an additional codon between the slippery sequence and the stop codon.

The other element required for the ribosomal frameshift is a secondary structure

downstream of the slippery sequence, which is referred to as the “pseudoknot” (Jacks et

al, 1988; Brierley1995). The pseudoknot structure has been suggested to contain two

stems. The stems 1 and 2 at the frameshift sites of EAV, PRRSV, and LDV, are

suggested to be 11-12 and 6- 7 nucleotides, respectively (den Boon et al, 1991; Godney et

16

Fig 1.3. Schematic representation of the life cycle of arteriviruses. Entering of

arteriviruses into susceptible cells occurs through receptor mediated endocytosis. At first

the viruses bind to specific receptors on the cell, and then the virus is taken into the cell

by clathrin-coated vesicles. The low pH stimulates fusion of virion envelope with the

endosomal membrane leading to the release of the viral nucleocapsid into the cytoplasm

of the infected cell. Translation and transcription are the next steps of the life cycle

followed by the replication of the genome and assembly of virion which happens by

budding the nucleocapsid via ER and Golgi complex. The final step is the exit of virion

which happens by exocytosis from the infected cells (adapted from Snijder and Spaan,

2007).

17

al, 1993; Meulenburge et al., 1993b). The frameshifting for EAV is reported to be 15% to

20% efficient (den Boon et al, 1991). Proteolytic processing of the polyprotein after

translation forms a structure to copy a full genome length complementary strand

(negative genome RNA) which functions as a replication template for the genome of

arteriviruses.

A nested series of subgenomic (sg) minus-strand RNAs (complementary of

sgmRNAs) are produced during transcription which will be discussed later. Each of (–)

sgRNA and (+) sgRNA has similar 5’ and 3’ ends (Snijder and Spaan, 2007). The

presence of specific structures for RNA replication of arteriviruses has yet to be

discovered. There are two NTRs located at the 5’ and 3’ ends of the genome of 156 to

221 and 59 to 117 nucleotides in lengths, respectively (Molenkamp et al., 2000a; Tijms et

al, 2001). The 3’ NTR has 33-47% identity in sequence between members of the

Arteriviridae family. A conserved sequence is found among all of these viruses in the 3’

NTR, located just before the poly A sequence (Godeny et al., 1993). In the 5’ end of

LDV, PRRSV, and SHFV, the start codon of ORF1a is located immediately after the

common leader sequence but for EAV, it is located 12 nucleotides downstream of the

leader sequence. This results in the presence of the entire genomic 5’ NTR in all of sg

mRNAs in LDV, PRRSV, and SHFV. There are interactions between host cell proteins

and the 3’ end of the negative-sense genomic RNA for SHFV, LDV, and EAV (Hwang

and Brinton, 1998). This indicates that for initiation of the (+) RNA transcription, the 3’

end of the (–) RNA and the interactions with the host proteins are crucial (Snijder and

Spaan, 2007). Using defective interfering RNAs of EAV, it has been found that the

presence of a 300 nucleotide region at both ends of the genome is required for efficient

18

replication. This suggests that NTR’s of both termini and some regions of the coding

sequence may contain signals required for replication of EAV (Molenkamp et al., 2000b;

Tijms et al, 2001). Similarly, interaction between the RNA hairpin secondary structure in

the 3’ NTR and the N protein gene, which is called the “kissing interaction”, has been

reported for PRRSV as vital for synthesis of viral RNA (Verheije et al., 2002).

Nidovirus transcription for generation of a nested set of sgmRNAs is a unique

feature of the replication cycle of these viruses. sgmRNAs are synthesized to express the

structural genes located at the 3’ end of the genome of these viruses (Baric et al., 1983;

Spaan et al., 1983; van Berlo et al., 1982, 1986; de Vries et al., 1990, 1997; Kuo et al.,

1991; Meulenberg et al., 1993b). The mRNAs of arteriviruses have common 3’ terminal

and 5’ leader sequences (Snijder and Horzinek, 1993; de Vries et al., 1997). The 5’ leader

sequence of sgmRNAs is the 5’ end sequence of the genome. The process by which this

leader sequence is merged into the sgmRNAs is the common mechanism of transcription

for Arteriviridae and Coronaviridae and is called “discontinuous transcription” (Snijder

and Meulenberg, 1998). The presence of conserved sequences at the “leader-body”

junctions of sg mRNAs of arteriviruses has been reported (de Vries et al., 1990; Chen et

al., 1993; Meulenberg et al., 1993a; Zeng et al.,1995; den Boon et al., 1996; Godeny et

al., 1998). A series of negative-strand (-) sg replicative intermediates (RIs) of EAV has

been detected. These (-)sgRIs, which are complementary of (+)sg mRNAs, are thought to

be involved in mRNA synthesis of arteriviruses (den Boon et al., 1996; Chen et al.,

1994). Fig. 1.4 is an illustration of the nested set composition of genomic and (+) and (-)

subgenomic strands of RNAs in Arteriviridae, and two proposed models for the

transcription are depicted. As demonstrated in Fig. 1.1 b and 1.4 a, the sg mRNAs of

19

Fig. 1.4. The nested set of RNAs in arteriviruses and the discontinuous strategy for the

mRNA transcription. (a) The structure of arteriviruses nested series of RNA. The positive

and negative strand RNAs are shown in white and black respectively. Below the full

strands of genomic and its complementary full RNA, two examples of + and – strand

subgenomic (sg) RNAs are depicted. The small white and black boxes on the RNAs

indicate the (-) and (+) transcription-regulating sequences (TRSs) respectively. The

leader and anti leader sequences are shown as hatched boxes. The sgmRNAs contain a

leader sequence drived from the 5’ end of the genome which is connected to the specific

sequences from 3’ end of the genome through a discontinouse mechanism of RNA

transcription. (b) and (c) show two proposed mechanisms for the transcription of

arteriviruses. The initial model for transcription was the leader-jump. In this method plus

strand RNA synthesis happens by falling the + leader sequence and base-pairing with the

– TRS on the downstream of the (-) genome RNA (b). However by the discovery of the –

sg RNAs the second model as body-jump (c) was suggested. In the body-jump method,

the body TRS falls during the synthesis of the –RNA. Then it base pairs with the leader

sequence leading to the synthesis of the – sg RNA which can be used as template for the

+sg mRNAs synthesis. The second method is more favorable by researches (Adapted

from Snijder and Meulenberg, 1998).

20

Arteriviridae contain the “leader” and “body” sequences. These two parts of the sg

mRNAs are not located in a continuous position in the genome sequence rather they are

transcribed from the 3’ and 5’-termini of the genome. Conserved sequences called

transcription regulating sequences (TRSs) connect the leader and body sequences of each

mRNA. The TRSs are located at the 3’ end of the leader sequence as well as at the 5’ end

of the body of each mRNA (Snijder and Meulenberg, 1998).

Two models have been proposed for mRNA transcription of members of the order

Nidovirales. The first model is called “leader-primed transcription” in which the

synthesis of the (+) sg RNA is disrupted and the (+) TRS located at the 3’ end of the

leader sequence complements with the (-) TRS of the 5’ end of each transcription unit

(Fig. 1.4b; Baric et al., 1983; Spaan et al., 1983). After the (+) and (-) TRS base-pairing

starts, it moves toward extension of the leader resulting in a sg mRNA. This model leads

to continuous synthesis of the (+) genomic RNA and discontinuous synthesis of the (+)

sg RNA (Snijder and Meulenberg, 1998). However, subsequent studies showed the

presence of a nested series of RIs that included (-) sg RNAs containing an anti-leader

sequence, and another model was proposed for RNA transcription. This model involves

discontinuous synthesis of (-) strand RNAs from the genome template (Fig. 1.4c). Base-

pairing of (-) leader TRS and (+) TRS of the template is crucial in both transcription

models (Snijder and Spaan, 2007). The TRS sequence for all members of the

Arteriviridae family has been determined. The sequence of TRS in the EAV genome is

highly conserved (den Boon et al., 1991). However, the TRS sequence is not the sole

determinant by which the virus transcription machinery chooses the correct TRS

sequence as the interaction of viral and/or cellular proteins, and viral RNA has also been

21

suggested to be involved in this process (Snijder and Spaan, 2007; Pasternak et al., 2004).

In addition to the leader TRS, 17 other conserved TRS sequences are detected in the

genome of EAV, but six of them are known to play a role in the EAV transcription

(Snijder and Meulenberg, 1998). The 3’ side of TRS is more conserved than the 5’ side.

This may indicate that the 3’ side of TRS is more important than the 5’ side (Chen et al.,

1993; Meulenberg et al., 1993a; Godeny et al., 1998). This finding suggests that “the

leader-primed model” is less favorable for viral mRNA transcription has been more

widely accepted in recent years.

3.4 Virion Assembly and Release from the Cell. The subcellular localization of EAV N

has been reported (Tijms et al., 2002). Since there is no evidence that N protein is

required for genome replication and transcription of arteriviruses, this protein has been

suggested to be involved in virus assembly. Viruses of the Arteriviridae family replicate

in the cytoplasm of host cells and their assembled nucleocapsid buds from the

intracellular membranes where the viral membrane proteins are already synthesized and

located (Fig. 1.3). The preformed nucleocapsid buds through the smooth endoplasmic

reticulum followed by the Golgi system. Then, it is released from cells by exocytosis and

finally by lysis of infected cells (Magnusson et al., 1970; Wood et al., 1970;

Stueckemann et al., 1982; Pol and Wagenaar, 1992).

4. Pathogenesis of Arteriviruses

4.1. Natural Infection and Persistence

4.1.1 EAV. EAV infection occurs via the respiratory route which is followed by virus

replication in alveolar macrophages and endothelial cells. The virus then tranclocates to

22

lymph nodes, and through the lymphatic system, it can spread to other host tissues.

Viremia is developed 3 days post-infection and virus can be isolated from all tissues by

this time (Balasuria and Maclachlan, 2004). Infection of horses with EAV may remain

subclinical, but clinical signs can include flu-like signs, abortion in pregnant mares, and

interstitial pneumonia in newborns (Bryans et al 1957; Balasuria and Maclachlan, 2004).

Injuries of the endothelial system and small muscular arteries, along with high

permeability of the vascular system are some of the mechanisms for pathogenesis of

EAV (Balasuria and Maclachlan, 2004). The EAV infection naturally results in persistent

infection affecting about 35% of infected male horses (Timoney and McCollum, 1993).

These “carrier stallions” are persistently infected animals that keep the virus in the

reproductive tract and shed it into the semen which helps to spread the virus (Snijder and

Spaan, 2007). The exact mechanism of the persistence of EAV is not fully understood.

4.1.2 PRRSV. Infection of pigs with PRRSV occurs via the respiratory route and it is

also transmissible to the fetus across the placenta (Zimmerman et al, 1997). The primary

cells that become infected are alveolar macrophages and, subsequently, macrophages in

other tissues also become infected. Viremia is seen at 12 to 24 hours post-infection and

may last 1 to 2 weeks in older pigs and up to eight weeks in piglets (Snijder and Spaan,

2007). Clinical features of PRRSV infection include high temperature, anorexia, lesions

in the lung, as well as stillborn, dead or weak piglets due to the reproductive problems of

infected sows. Different factors influence the outcome of infection with PRRSV, which

include the age, gender, and immune condition of the host and the strain of virus

infecting the host (Wensvoort et al., 1993).

23

4.1.3 LDV. Infection of the host by LDV is asymptomatic but results in a permanent

viremia in the infected animals. Maintenance of LDV happens by constant replication of

the virus in a small group of peritoneal macrophages (Onyekaba et al., 1989b). The titer

of virus reaches 1010

lethal dose 50% (LD50) per ml of plasma at 24 hours post-infection,

but afterwards it declines to 104

ID50 to 106 ID50. The latter is the titer of LDV which

persists along with the increased amount of lactate dehydrogenase through the life of the

infected mouse. Virus can be detected in spleen, lymph nodes, thymus, and liver of

infected mice during the persistent period (Cafruny et al., 1986). In specific inbred lines

of mice, a deadly poliomyelitis resulting from infection by neurovirulent strains of LDV

can occur, which is age-related (Contag and Plagemann, 1989).

4.1.4 SHFV. SHFV leads to asymptomatic, acute or persistent infections in African

monkeys based on the strain of the virus which infects these animals (Gravell et al.,

1986a; London, 1977). However, the virus caused deadly hemorrhagic fever outbreaks in

captive macaques kept in the National Institutes of Health (NIH) quarantine which were

shipped from India (Tauraso et al., 1968). Pathogenesis of SHFV in macaques is largely

unknown. Fever, erythem face, edema, dehydration, and hemorrhages are the

manifestations of SHFV infection in macaques. Normally, the SHFV-infected macaques

die by two weeks post-infection and the rate of mortality is near 100%. The target cells

for SHFV in the host are macrophages, similar to other members of the family (Gravell et

al., 1986b).

4.2 Immune Response to Arteriviruses Infections.

4.2.1 Innate Immune Response to Arteriviruses. Characteristics of innate immunity to

arteriviruses include production and expression of pro-inflammatory cytokines and

24

interleukins, and activation of infiltrating natural killer (NK) cells and macrophages

(Snijder and Spaan, 2007). Of the pro-inflammatory cytokines, induction of tumor

necrosis factor-alpha (TNF-α) has been reported in vitro in macrophages infected with

the virulent and avirulent strains of EAV. In the case of infection by avirulent EAV

strains, production of TNF-α was lower (Balasuriya and Maclachlan, 2004; Moore et al.,

2003). The induction of innate immune mechanisms is believed to be poor in PRRSV

infection (Murtaugh et al., 2002). Lee et al. (2004) demonstrated that induction and

sensitivity to interferon-alpha (IFN-α) in porcine alveolar macrophages varies among

different field isolates of PRRSV. PRRSV was shown to suppress the activation of type I

IFN transcription factor, interferon regulatory factor 3 (IRF3), in MARC-145 infected

cells and it is suggested that PRRSV N is involved in the deactivation of IRF3 (Lai F.W.,

2006). Several studies indicate that non-structural (nsp) proteins of PRRSV have

important role in suppression of innate immune response. For example two auto-clevage

products of nsp1 (nsp1-alpha and nsp1-beta) are involved in inhibition of synthesis and

signaling pathway by IFN-beta (Chen et al., 2009a). Moreover, in cells expressing two

nsp1-alpha, nsp1-beta, and nsp11 of PRRSV, double-stranded RNA (dsRNA) signaling

pathway was inhibited. The correlation of nsp1-beta to inhibit gene induction by IRF3

and NF-kappaB mediated by dsRNA and Sendai virus is reported (Beura et al., 2010).

Also the down-regulated expression of IL-1 beta and TNF-alpha in cells infected with

virus carrying the mutant forms of nsp2, suggests a regulatory role for this protein in

innate immue respone for PRRSV (Chen et al., 2009b). LDV has been reported to

activate NK cells in the infected host, which results in elevated interferon-gamma (IFN-γ)

25

in the serum, although elevated levels of this cytokine cannot clear the infection

(Markine-Goriaynoff et al., 2002).

4.2.2 Adaptive Immune Response to Arteriviruses. As mentioned above, persistent

infection is a common feature of arteriviruses. It can be as long as 2-3 months in the case

of PRRSV-infected swine, or may be a life-long in LDV-, EAV-, and SHFV-infected

hosts (1992; Onyekaba et al., 1989a; Timoney and McCollum, 1993; Gravell et al.,

1986b). The mechanism of evasion from the host immune system by arteriviruses is not

known.

The involvement of antibody-mediated immune response in arterivirus infection

has been studied for different members of the family. One week post-infection, antibodies

against viral proteins can be detected (Coutelier et al., 1986; Gravell et al., 1986b;

McCollum, 1986; Nelson et al., 1994). In EAV-infected animals, antibodies are induced

against N, M, GP5, and GP2 proteins (Balasuriya et al., 2002; MacLachlan et al., 1998).

In PRRSV-infected pigs, serum antibodies are present against GP2 to GP5, N, and M. In

the serum of these animals, the most abundant antibody is against the N protein (Loemba

et al., 1996; Meulenberg et al., 1995b).

Most of the neutralizing antibodies in animals infected with arteriviruses are

induced against GP5 (Cafruny et al., 1986; Chirnside et al., 1995a, 1995b; Gonin et al.,

1999). In EAV-, LDV-, and PRRSV-infected animals, the presence of neutralizing

antibodies against GP5 has been reported with the neutralization epitopes located in the

ectodomain of GP5 (Balasuriya et al., 2004; Chirnside et al., 1995 a, 1995b; Li et al.,

1998; Plagemann, 2004). The ectodomain of GP4 of PRRSV seems to harbor the binding

domain for a second MAb (Meulenberg et al., 1997). Virus neutralization by MAbs

26

raised against GP5 is more efficient than by MAbs against GP4 (Wieland et al., 1999).

The immune response against structural proteins of SHFV has not been investigated

thoroughly (Snijder and Spaan, 2007).

In EAV-infected horses, highly neutralizing antibodies appear at the same time as

the virus is cleared. These antibodies remain in the host for a long time and they have

been suggested to be important for protection against EAV (Snijder and Meulenberg,

1998). Low titers of neutralizing antibodies against LDV and PRRSV have been

observed and it seems that these antibodies do not clear or lower virus burden in the

blood (Cafruny et al., 1986; Loemba et al., 1996; Yoon et al., 1996). It has been reported

that antibodies against LDV and PRRSV affect virus intake by macrophages in vitro.

This suggests that the complex of virus-antibody in infected mice and pigs may increase

the infection (Cafruny and Plagemann, 1982; Yoon et al., 1996). It has been shown that

maternal antibodies can protect progeny of LDV- and PRRSV-infected animals against

these infections (Broen and Cafruny, 1993; Snijder and Meulenberg, 1998). Antibodies

against LDV can inhibit the progression of infection towards age-dependant poliomyelitis

in mice (Harty et al., 1987; Harty and Plagemann, 1990; Plagemann and Moennig, 1992).

The factors involved in LDV persistence include infection of newly developed

macrophages leading to slow and consistent formation of virus-susceptible cells,

emergence of immune resistant virus variants during the persistent infection, and

presence of infectious virus-antibody immune complexes in LDV infected mice

(Onyekaba et al., 1989a; Rowland et al., 1994; Monteyne and Coutelier, 1994; Chen et

al., 1997). The antibody-mediated response against SHFV is dependent on the species of

the infected monkeys as well as virus variants. There is no efficient immune response in

27

infected macaque monkeys which die promptly after SHFV infection. However, in

infected Patas monkeys, two different antibody responses have been shown, depending

on SHFV strains causing different forms of the disease. The SHFV strains causing acute

infection induce high titer of neutralizing antibodies against the virus and this coincides

with the complete disappearance of virus from blood. However, the viral strains that

cause persistent infection of the patas monkeys induce only low titers of non-neutralizing

antibodies against the virus (Gravell et al., 1986b).

Cell-mediated immune response against Arteriviridae has not been well studied.

Cell-mediated immune (CMI) response seems to play an important role in the immune

response against SHFV. Subsequent clearance of super-infection in Patas monkeys which

had already been persistently infected with a different strain of SHFV is supportive of

CMI response to SHFV. Moreover the lack of cross- neutralization between the

antibodies raised against the two different strains of SHFV supports CMI involvement as

well (Gravell et al., 1986a, Snijder and Meulenberg, 1998). Experiments on EAV

infected ponies showed that cytotoxic CD8+T-cells are mediators of the cell-mediated

response against EAV and that response is specific for the virus (Balasuriya and

MacLachlan, 2004; Castillo-Olivares et al., 2003). The presence of cytotoxic T

lymphocytes (CTLs) and helper T cell responses stimulated by LDV infection has been

shown. These responses lasted for 30 and 250 days post-infection in different

investigations, but were not effective in prevention of LDV replication (Even et al., 1995;

van den Broek et al., 1997). Both CD4+ and CD8

+ responses have been detected 4 to 8

weeks post-PRRSV infection (Murtaugh et al., 2002; Xiao et al., 2004). T-cell

proliferation responses against M, GP2, and GP5 have been detected between 4-12 weeks

28

after PRRSV infection (Bautista et al., 1999). More investigations are needed to

determine if cell-mediated immunity is involved in protecting animals from infection

with arteriviruses.

5. Nuclear Localization of Viral Proteins in (+) ssRNA viruses

5.1 Nucleolar and Nuclear Localization of Proteins. The nucleolus is a part of

eukaryotic cell which is known for ribosome biogenesis (Melese and Xue, 1995; Sirri et

al., 2008). Transcription of rRNAs, cleavage of ribosomal RNA precursors, and

aggregation of ribosomal RNAs with ribosomal protein to construct the 40S and 60S

subunits of the ribosome all occur in the nucleolus (Sirri et al, 2008). Using electron

microscopy (EM), three distinct parts of the nucleolus have been observed. The “dense

fibrillar component” (DFC) and the “granular component” (GC) are two major

compartments of the nucleolus observed by conventional EM (Jordan 1984). The third

compartment is called the “fibrillar center” (FC) which comprises a small portion of the

nucleolus (Shaw and Jordan 1995). The FCs and DFCs are inside the GC which is

composed of granules that are 15-20 nm in size. When proteins are in nucleus, either

transferred by importins or diffused, some are localized to specific parts of the nucleus,

such as nucleoli, and others are seen all over the nucleus (Sirri at al, 2008; Andersen et al.

2002, 2005; Leung et al. 2003). All of the required proteins for ribosomal synthesis are

cycling between the nucleolus and the nucleoplasm in the interphase stage of the cell

cycle. Also, the diffusion of nucleolar proteins between these compartments of the

nucleus is a permanent feature of nucleus, which happens very rapidly (Dundr et al. 2004;

Sirri et al, 2008). Many factors are important in the nucleolar localization of a protein.

The most important factor for this activity is the localization of the protein to the nucleus

29

which is followed by other mechanisms that lead a protein to the nucleolus. The

mechanisms and signals for nuclear localization of a protein and their export to the

cytoplasm have been investigated thoroughly, but the molecular bases for nucleolar

localization of proteins are still to be discovered (Sirri et al, 2008). The cytoplasm and

nucleus are separated by the nuclear envelope (NE). The NE is a continuation of the

endoplasmic reticulum (ER). It contains many holes, named nuclear pore complexes

(NPCs), for transferring macromolecules between the nucleus and the cytoplasm. Some

of these macromolecules are imported from the cytoplasm including nuclear proteins

such as histones and transcription factors. Some molecules synthesized in the nucleus

must be exported to the cytoplasm, including transfer RNA (tRNA), ribosomal RNA

(rRNA), and messenger RNA (mRNA). The NPCs are the only path through which these

translocations happen. The numbers of NPCs differ depending on the size and activity of

each cell. For example, a human cell can contain 3,000-5,000 NPCs in the NE. NPCs

have octuple rotational symmetry and each of them contains a central core compartment

inserted in the NE with extended cytoplasmic and nuclear elongations to shape the

cytoplasmic extensions and the nuclear basket structures, respectively. The proteins

constituting the NPCs are called nucleoporins which also provide the place that nuclear

transport proteins and elements dock and interact for their activities (Gorlich and Kutay,

1999). NPCs shape the routes through which all nuclear trafficking processes occur

(Fedherr et al, 1984; Gorlich and Kutay, 1999). The trafficking of small molecules less

than 9 nm in diameter, such as metabolites, takes place by passive diffusion through these

channels (Paine et al, 1975; Gorlich and Kutay, 1999). On the other hand, the

translocation of larger macromolecules greater than ~40 kD through the NPCs happens

30

Fig. 1.5.Schematic illustration of nuclear protein transportation. The cargo protein

containing a nuclear localization signal (NLS) forms an import complex with importin-α

(α) and importin-β (β). This complex docks onto the nuclear pore complex (NPCs) and

enters into the nucleus where the binding of RanGTP to imp-β separates it from the

complex. This is followed by disassembly of the cargo and imp-α in the nucleus and

return of the imp-α to the cytoplasm by its nuclear export protein, CAS in association

with RanGTP. In the cytoplasm RanGAP induces the hydrolysis of GTP and releases

imp-α. (From Cook et al., 2007).

31

actively and is a signal-mediated process regulated by the presence of a particular signal

on the protein and its interaction with transportins and NPCs. This mechanism is also

involved in transferring some smaller proteins and RNAs into the nucleus (Breeuwer and

Goldfarb, 1990; Jäkel et al., 1999; Gorlich and Kutay, 1999, Stewart, 2007). The first

identification of nuclear translocation signal on a protein was reported for the

nucleoplasmin of Xenopus laevis oocytes (Dingwall et al 1982), and the simian virus 40

(SV40) large-T antigen was the first protein reported to contain the nuclear localization

signal (NLS) for import into the nucleus (Kalderon et al., 1984; Lanford and Butel, 1984;

Robbins et al., 1991; Gorlich and Kutay, 1999). Three types of NLSs have been reported

so far. “Pat 4” is a type that includes a stretch of four basic amino acids of mostly lysines

or arginines. “Pat 7” is a second type of NLS which contains a motif of seven amino

acids starting with a proline followed by two other amino acids which are followed by

four other amino acids and three of them being basic residues. The third type of NLS is

“bipartite” which begins with two basic amino acids followed by 10-12 residues as a

spacer and ends with five amino acids, among which at least three are basic (Robbins et

al., 1991; Nakai and Kanehisa,1992). It is now accepted that in NLS-related nuclear

import, four essential components are involved. Two of them are importin α (Impα),

importin β (Impβ), and two others are Ran and nuclear transport factor 2 (NTF2) which

belong to the RanGTPase system (Adam and Adam, 1994; Gorlich et al., 1994; Chi et al.,

1995; Gorlich et al., 1995; Melchior et al. 1993; Moore and Blobel, 1993; Moore and

Blobel, 1994; Paschal and Gerace, 1995). More investigation has shown that different

nuclear transfer pathways are involved in translocating proteins into the nucleus. Most of

these mechanisms include a super family of carrier proteins called β-karyopherins.

32

Among them are Impα and Impβ which mediate the nuclear import of many proteins.

Often the complex of Impβ-Impα transfers the protein of interest into the nucleus, in

which Impα acts as an adaptor for Impβ by recognizing the NLS on the cargo protein

(Stewart, 2007). Every minute, approximately 100-1000 cargo proteins are transported

through each NPC by the classical pathway of nuclear protein import. The import cycle

includes four phases: the cargo-carrier complex formation in the cytoplasm, transfer via

NPCs, dissociation of the import complex in the nucleus, and recovery of the importins

(Fig. 1.5). First in the cytoplasm, the cargo protein containing NLS binds to the Impβ-

Impα heterodimer via the NLS binding site on the Impα which is acting as an adaptor.

The import complex of cargo and Impβ-Impα-heterodimer then moves towards the NPCs

and docks there. By Ran-GDP, this complex passes via the NPCs. In the nucleus, the

present Ran-GTP binds to Impβ resulting in disassembly of the import complex and

liberation of the cargo protein into the nucleus in which it either diffuses to a specific site

or spreads evenly. The last step is recycling importins and RanGTP. Impβ returns to the

cytoplasm along with RanGTP and Impα leaves the nucleus along with the β-karyopherin

CAS (Cellular Apoptosis Susceptibility protein) and RanGTP. In the cytoplasm, the Ran

GTPase activating protein (RanGAP) prompts the Ran GTPase to produce RanGDP

leading to release of the importins (Stewart, 2007). Protein import is an orchestrated

cycle which involves groups of controlled interactions between cargoes and carriers.

These interactions are coordinated by the nucleotide condition of Ran which is cycling

between GTP- and GDP-bound forms of Ran controlled by its Guanine nucleotide-

exchange factor (RanGEF) and RanGAP (Ran GTPase activating protein). The RanGEF

resides in the nucleus and catalyses the recharging of RanGDP with GTP. RanGAP is

33

cytoplasmic and stimulates hydrolysis of GTP. The RanGTP is localized in the nucleus

and RanGAP is in the cytoplasm. The positioning of RanGTP in the nucleus mediates

export of Impβ protein from the nucleus to cytoplasm where GTP is hydrolysed leading

to the release of RanGDP to the cytoplasm preparing to be used in another cycle for

protein import. In addition to the nucleotide state of Ran, its conformational changes

during the nuclear import process are also determinants of its interaction with the

importin family members and the energy produced by RanGTPase for nuclear import

(Stewart, 2007). Hydrolysis of RanGTP in the cytoplasm and exchange of the nucleotide

state of Ran from GTP to GDP in the cytoplasm retains a gradient of RanGTP/RanGDP

which is supported by the presence of the RanGAP in the cytoplasm and RanGEF in the

nucleus. This gradient determines the direction of nuclear transport (Cook et al., 2007).

5.2 Nucleolar Localization of (+)ssRNA Virus Proteins. Many viruses including RNA

viruses, DNA viruses, and retroviruses have molecular interactions with the nucleus and

nucleolus. The nucleus is the place in which many DNA viruses, retroviruses, and a few

RNA viruses replicate. Positive-strand RNA viruses mostly replicate in the cytoplasm of

infected cells and do not require the function of the nucleus for their replication cycle.

Their essential proteins have been shown to localize in the nucleus which is not the place

for their replication (Hiscox, 2007; Sirri et al, 2008; Wilhelmsen, et al., 1981; Evans et

al., 1980). Nucleolar localization of many proteins of positive-strand RNA viruses,

particularly the nucleocapsid protein, has been a topic of interest for many researchers.

The nucleocapsid protein of coronaviruses such as IBV (infectious bronchitis virus),

SARS (severe acute respiratory syndrome) virus, and MHV (mouse hepatitis virus), the

capsid protein and nonstructural protein nsP2 of alphavirus, and the core protein of

34

dengue virus, are all examples of (+)ss RNA viruses which have been reported to localize

to the nucleus and nucleolus of infected cells (Hiscox et al 2001; Wurm et al., 2001; You

et al. 2005; Timani et al., 2005; Tijms et al., 2002; Rowland et al., 1999; Michel et al.,

1990; Rikkonen et al., 1992; Wang et al., 2002). In order to fulfill their cytoplasmic

activities such as their involvement in the structure and replication of viruses, these viral

proteins must translocate back from the nucleus to the cytoplasm. It is well known that

the presence of an NLS motif is a vital part of nuclear transport for many viral proteins.

However, the nucleolar localization signal (NoLS) is not well characterized for viral or

host proteins (Hiscox, 2007). In some viruses, NLS and NoLS work in harmony to

localize a protein to the nucleus. The presence of NoLS with a functional NLS motif in

the sequence and resemblance to host NoLSs has been well studied for PRRSV

nucleocapsid protein. Two NLS motifs have been identified in PRRSV N: the ‘pat4’ type

NLS-1 at positions 10-13 and the ‘pat7’ type NLS-2 located at 41-47. The cooperation of

NLS and NoLS in the case of PRRSV N results in translocation of this protein into the

nucleolus. By constructing truncated proteins fused to enhanced green fluorescent protein

(EGFP), the region of PRRSV N responsible for nucleolar localization has been mapped

to position 41-72 which contains the functional NLS-2 motif. The N protein also contains

a leucine rich motif located at 106-117, which is the nuclear export signal (NES)

responsible for transferring the protein back to the cytoplasm (Rowland et al., 2003;

Rowland and Yoo, 2003). The other way to transport a protein to the nucleolus is

through collaboration of the viral protein with cellular proteins. The presence of a

proposed NoLS in the hepatitis delta virus (HDV) delta antigen is responsible for both

nucleolar localization and binding to the nucleolar protein nucleolin. Mutation of the

35

NoLS of HDV delta antigen prevents this protein from either translocating into the

nucleus or binding to nucleolin. This implies that nucleolar translocation is correlated

with binding to the host protein nucleolin (Lee et al, 1998). Unlike NLS, there is no

available program to predict NoLS. Investigation on the IBV N protein nucleolar

localization has revealed that a motif of 8 basic amino acids on this protein functions as a

NoLS signal and indeed interacts with nucleolar protein leading the IBV N protein into

the nucleolus. This motif is also an interactive part of IBV N with nucleolar proteins

(Reed et al., 2006; Hiscox, 2007; Sirri et al, 2008).

5.3 Function of Nuclear Localization of Viral Nucleocapsid Proteins. The presence of

NoLS on viral proteins indicates that viruses use the nucleolus for specific reasons and

suggests that viruses have undergone evolution to manifest and maintain this function.

Evidence suggests that the nuclear or nucleolar localization of some (+) SS RNA viruses

influences the pathogenesis and virulence of these viruses. Examples of viral proteins

affecting the virus replication and pathogenesis are explained below.

The non-structural protein nsP2 of Semliki Forest virus has been shown to

localize to the nucleus and by changing one amino acid in this protein, its trafficking to

the nucleus was interfered, which resulted in lowered virulence of the virus (Peranen et

al., 1990; Fazakerley et al., 2002). Lee et al., (2006b) by infecting pigs with an infectious

clone of PRRSV which had mutations in its functional pat7 NLS, demonstrated that

viremia was shorter. This was the first in vivo report of the importance of nuclear

localization in the pathogenesis of (+)ssRNA viruses (Lee et al., 2006a, 2006b). Another

example is the core protein of Japanese encephalitis virus (JEV) which has been reported

to improve the replication of JEV. By eliminating the nuclear and nucleolar localization

36

activity of JEV core protein, the recombinant virus showed reduced replication compared

to the wild-type virus (Mori et al., 2005). Interaction of the JEV core protein with the

nucleolar protein B23 has been shown (Tsuda et al., 2006). Moreover, the newly

synthesized viral RNA was found in the nucleus along with the viral replicase in infected

cells. Nucleolar localization of the flavivirus core protein has also been suggested to be

important in viral RNA synthesis (Uchil et al., 2006). Interaction of many (+) ssRNA

viruses with the nucleus and nucleolus is believed to help viral replication.

Another example of RNA viruses that interact with nucleus is the retroviruses.

The best studied example is human immunodeficiency virus (HIV). The genome of HIV

includes two copies of positive-sense RNA and its replication takes place in two separate

phases: cytoplasm and nucleus of infected cells. In the cytoplasmic phase, the viral

genome is first reverse-transcribed by the viral RT and then transferred to the nucleus

where the viral genome is transcribed and is transported back to the cytoplasm (Hiscox,

2007).

One of the first assumptions for the nucleolar localization of capsid and other

viral proteins of RNA viruses was that they diffuse through NPCs to reside in the nucleus

without any impact on the virus life cycle. But the possible selective pressure on keeping

this phenomenon during RNA virus replication leads to the idea that this function is a

helpful strategy for efficient viral replication. This maybe achieved by occupying the

nucleolus of the cell and using the nucleolar proteins involved in cellular transcription

and ribosomal biogenesis for virus benefit (Hiscox, 2007). Disruption of the nucleus

functions seems to be one of the influences of nucleolar localization of viral proteins.

Such an effect can be related to the reduced or elevated levels of the nucleolar proteins in

37

the nucleus, nucleolus, and cytoplasm as a result of moving or reorganization of these

proteins by viral proteins (Hiscox, 2002). The result of nucleolar localization of viral

proteins could interfere with the structure and activities of the nucleolus in virus-infected

cells. For instance, the disruption of the nucleolar functions has been reported in

poliovirus-infected cells. In poliovirus-infected cells, nucleolin is relocated to the

cytoplasm from the nucleolus (Waggoner and Sarnow 1998). Also, inactivation of the

upstream binding factor (UBF) has been reported for poliovirus which leads to the

inhibition of the RNA polymerase I-mediated transcription in virus-infected cells

(Banerjee et al., 2005). The nucleolar structure has been shown to be disrupted in IBV-

infected cells (Dove et al., 2006a). Moreover, G2 arrest of the cell cycle and abnormal

cytokinesis have been reported in IBV-infected cells (Wurm et al., 2001; Dove et al.,

2006b). The nucleocapsid proteins of IBV and PRRSV have been shown to interact with

nucleolar proteins, nucleolin and fibrillarin (Chen et al., 2002; Yoo et al., 2003). HIV-I

Rev protein is another example of a viral protein from (+) ssRNA viruses is located in the

DFCs and GCs compartments of the nucleolus and disrupts the architecture of nucleolus

and interacts with B23.1 (Miyazaki et al., 1996). The high number of cells at the G2 and

M stages of cell cycle (Miyazaki et al., 1996) and failed cytokinesis (Cannavo et al.,

2001) are two changes in Rev expressing cells.

It has been suggested that RNA viruses hijack the nucleolus by recruiting its

proteins during their replication. These nucleolar proteins have vital functions in

synthesis, processing and translation of cellular RNA. For example, picornaviruses

interact with and use nucleolin during their replication. Nucleolin interacts with the

picornavirus internal ribosome entry site (IRES) located at the 5’ end of its genome. This

38

Fig 1.6. Schematic presentation of a normal cell cycle. The cell cycle has two major

divisions: mitosis (M) and interphase. The second starts after mitosis and include three

phases: G1 which is the stage of preparing cells for the next stage the S phase in which

the DNA synthesis happens and is followed by G2 phase which prepares cells for the

next M. Each stage is regulated by different cyclines and Cdks (adapted from Schafer,

1998).

39

induces the IRES mediated translation which is cap-independant (Izumi et al., 2001).

Nucleolin also has been shown to interact with the 3’ non-coding terminus of the genome

of picornaviruses. This region acts as a regulator of the negative-strand RNA synthesis at

the early stages of poliovirus infection (Waggoner and Sarnow, 1998). Nucleolin

interacts with IRES in hepatitis C virus (HCV) and its interaction with HCV NS5B

protein has been proposed to be important in NS5B function (Hirano et al. 2003).

Localization of viral proteins in the nucleolus can abolish the translation process in

infected cells. Investigations have shown that nucleolar localization of the precursors of

3C protease of human rhinovirus, 3CD` and 3CD, at the beginning of the infection

prevents cellular RNA transcription by their protease activity (Amineva et al., 2004).

Such a shut-down of cellular protein translation has been reported as a result of nucleolar

localization of encephalomyocarditis virus (EMCV) 2A protein (Aminev et al, 2003).

5.4 Modulation of the Cell Cycle by Viral Proteins. The cell cycle is a synchronized

and controlled process for the growth and extension of cells. It is involved in the growth

and development of the organism as well as other biological functions such as repairing

DNA damage, tissue growth in injuries and cancer development. This process is

regulated by several proteins which orchestrate a series of phases leading to the formation

of two daughter cells. The key regulatory proteins for the cell cycle include the cyclin-

dependent kinases (Cdks) and the cyclin proteins which control moving cells through

four phases: G1 (gap 1), S (synthesis), G2 (gap 2), and M (mitosis) (Fig. 1.6).

Morphological subdivisions of cell cycle include two distinct phases, interphase and

mitosis (M). The interphase includes the G1, S, and G2 phases of cell cycle and the M

phase comprises of four stages of prophase, metaphase, anaphase, and telophase.

40

Interphase and M are two clear stages of cell cycle involved in DNA synthesis and

division of cells into two daughter cells, respectively. These two are separated by “gaps”

called G1 and G2. In the G1 step, the cells provide enzymes and proteins involved in

DNA synthesis which happens in the S phase, followed by the G2 in which the cell

becomes ready for M phase. Therefore, the DNA content of the cell at the G1, S, and G2

are 2N, 4N, and 4N, respectively. G0 is the quiescent stage during which cells are not

active and do not cycle (Schafer, 1998).

The key regulatory factors of cell cycle are cyclin proteins and Cdks. Cdks are a

group of serine/ threonine kinases and are vital for the progression of cell cycle. Cdks

become activated mostly by phosphorylation of their threonine and tyrosine residues.

Each stage of cell cycle contains specific regulatory factors which are neither active nor

abundant in the other phases of the cell cycle (Schafer, 1998). Many pathogens including

viruses have been reported to modulate the cell cycle. Viruses intervene with the cell

cycle by different strategies including: destroying the host cell normal growth to replicate

more efficiently, providing a better situation for virus genome replication, and prolonging

the cell cycle at different phases to provide more time for virus assembly. Most of

investigations of viral effects on cell cycle have been for DNA viruses and retroviruses.

However, interest in the interference of RNA viruses with cell cycle has been increasing.

Cell cycle control by cyclines and CdKs has been studied intensively, though recent

investigations show that nucleolus has direct or indirect effects on cell cycle along with

the aforementioned regulatory factors. Interaction of RNA viruses and their proteins with

nucleolus has been of great interest for researchers in this area (Emmett et al., 2004).

There are two major mechanisms that DNA viruses use to interfere with cell cycle. The

41

first strategy is used largely by DNA viruses with a large genome, such as herpesviruses,

which stops cells at the G1 stage inhibiting their transfer to the S phase of cell cycle

(Flemington, 2001). The second strategy is mostly used by DNA viruses with a small

genome, such as adenoviruses, which encode proteins interfering with regulatory factors

of the cell cycle (Moran, 1993). Cell cycle interruption has been well studied for

retroviruses. HIV infection leads to accumulation of infected cells in the G2 phase of cell

cycle which increases virus propagation. The viral protein responsible for this

phenomenon has been identified as Vpr, which localizes to the nucleus (Braaten et al,

1995; Poon et al, 1998). Negative- and positive-stranded RNA viruses have been reported

to modify cell cycle at different stages. For instance, the G0 arrest of cell cycle in cells

infected with measles virus, a negative ssRNA virus, has been reported (Naniche et al.,

1999), and the paramyxovirus simian virus V protein has been shown to slow cell cycle at

G1 and G2 stages (Lin and Lamb, 2000). Cell cycle modifications by positive-sense

RNA viruses have also been investigated. IBV accumulates infected cells at the S/G2

stage and inhibits normal cytokinesis (Chen et al., 2002; Wurm et al., 2001). Replication

of coxsackievirus is increased in cells arrested in G1 or G1/S phases (Feuer et al., 2002).

The other mechanism by which viruses interact with cell cycle control is by targeting

sub-nuclear structures involved in cell cycle such as nucleoli and their associated proteins

(Hiscox, 2002; Hiscox 2003). The concentration and distribution of nucleolar proteins

such as nucleolin, B23, and fibrillarin have been shown to be involved in regulation of

the cell cycle process (Sirri et al., 1997; Azum-Gelade et al, 1994; Fomproix et al., 1998).

Investigations have indicated that both non-structural and structural proteins of different

viruses can influence cell cycle progression. Among non-structural proteins, the

42

following have been implicated: the Vpr protein of HIV-1 which arrest the cell cycle in

G2/M (Mahalingam et al, 1997; Matsuda et al, 2003); σ1S protein of alpha reovirus

stalls cells at the G2/M phase (Poggioli et al, 2000); the V protein of the paramyxovirus

simian virus 5 (SV5) slows the cell cycle progress (Lin and Lamb, 2000); the non-

structural proteins 5A and 2 of hepatitis C virus regulate cell cycle phases (Ghosh et al,

1999; Yang et al, 2006), and MHV NS protein p28 and SARS 7a protein halt the cell

cycle at G0/G1 (Chen et al, 2004, Yuan et al, 2006). Among the structural proteins of

viruses, studies indicate that transmissible gastroenteritis virus nucleocapsid protein

(TGEV N) and IBV N play a role in cell cycle regulation. TGEV N is suggested to

modify cell cycle by increasing the number of dividing cells (Wurm et al, 2001) and the

nuclear localization of IBV N has been shown to be influenced by the cell cycle phase. In

synchronized Vero cells at the G2/M phase of the cell cycle upon transfection by IBV N

protein, the number of cells which showed nuclear localization for the N protein were

increased (Cawood et al, 2007).

5.5 Nucleocapsid Protein of Arteriviridae. The nucleocapsid (N) protein is the most

abundant protein in the arterivirus particle, comprising up to 40% of the virion protein.

The N protein is encoded by the 3’ most ORF of the genome. It is a basic protein and has

a molecular weight of 12-15 kDa (de Vries et al., 1992; Faaberg and Plagemann, 1995;

Godeny et al., 1995; Bautista et al., 1996). EAV N and PRRSV N are phosphorylated

(Zeegers et al., 1976; Wootton et al, 2002). The N protein is present as disulfide-linked

homodimers in the PRRSV particles (Mardassi et al., 1996; Meulenberg and Petersen-den

Besten, 1996) but the N of EAV and LDV has been reported to be in a monomeric form

in viral particles (de Vries et al., 1995a; Faaberg et al., 1995a). The N protein is an RNA

43

binding protein and the RNA binding domain for PRRSV N has been shown to overlap

with the functional NLS of this protein (Rowland and Yoo, 2003). In the sera from

infected mice with LDV, the level of antibodies against N is very low (Coutelier et al.,

1986; Faaberg and Plagemann, 1995). But in infected animals with EAV and PRRSV,

antibodies against N are abundant, which makes the N protein a good candidate for

serodiagnosis (Chirnside et al., 1995c; Kheyar et al., 1997; Meulenberg et al., 1995a;

Denac et al., 1997; Rodriguez et al., 1997). The N proteins of EAV and PRRSV have

been shown to localize to the nucleus and nucleolus of virus-infected and gene-

transfected cells (Tijms et al., 2002; Rowland et al., 1999). The PRRSV N has been

shown to interact and colocalize with the nucleolar protein fibrillarin, bind to the viral

and ribosomal RNA, and partner with the inhibitor of MyoD family-a (I-mfa) domain-

containing protein (HIC) (Yoo et al., 2003; Song et al., 2009). It has been shown that HIC

interacts with a T1 type of cyclin which in turn is part of the transcription system and is

involved in cell cycle progression (Young et al., 2003). Further studies have shown that

PRRSV infection using a mutant virus with disabled NLS is clinically attenuated

compared to the wild-type virus, indicating the importance of nuclear localization in the

viral pathogenesis (Lee et al., 2006a, 2006b; Pei et al., 2008). The interaction and

colocalization of N with the nucleolar proteins involved in cellular activities, such as cell

cycle, suggests that N has a regulatory function to influence the cell cycle process, in

addition to the structural function for virion assembly.

44

II.OBJECTIVES OF THE THESIS

The nucleolar localization of N protein of arteriviruses demands further

investigation for its biological function and whether it is a common feature of this family

in the Nidovirales order. The interactions of PRRSV N with the nucleolar and cellular

proteins result in modulation of cellular activities. Previous studies of coronavirus N

proteins and their modulation of cell cycle provides a basis for the hypothesis that the

capsid protein of Arteriviridae has a major role in cell cycle progression during virus

replication. Thus, the main goal of this study was to elucidate the nucleolar localization

of capsid proteins of all four member viruses of the Arteriviridae family and involvement

in the cell cycle.

Outline of Specific Objectives. Investigation on N protein has been mainly

accomplished using PRRSV, and information on other member viruses within the family

Arteriviridae is limited. Therefore, the first objective of this study was to investigate the

nuclear and nucleolar localization activity for N proteins of LDV, EAV, and SHFV.

The N protein of LDV was cloned as a GFP fusion protein and examined for its

subcellular distribution. The LDV N nucleolar localization signal was identified and

structure-function studies were conducted using mutant constructs. Subsequently, key

amino acids for the nuclear localization of LDV N were determined. Based on

experimental data and information from PRRSV N, the conclusion was made that LDV N

uses an active nuclear transport mechanism (Chapter 2).

Following the study of LDV N, our investigation was extended to EAV N and

SHFV N proteins. Using EGFP-EAV N and EGFP-SHFV N genes, subcellular

localization of the EAV N and SHFN explored. A functional NLS was identified for

45

EAV N and SHFV N and mapped using a series of mutant constructs. This was followed

by investigating of the molecular basis for their nuclear localization which (Chapter 3).

PRRSV N interacts with nucleolar and cellular proteins which play a role in

regulating cell cycle. This led us to hypothesize that the capsid protein of Arteriviridae

has a regulatory role in the cell cycle during virus replication. Therefore, in Chapter 4,

cell cycle modification by arteriviruses was investigated using PRRSV and EAV in virus-

infected cells. The role of N in cell cycle modification was then studied using N genes of

all 4 member viruses within the family by over expression of N in gene transfected BHK-

21 cells (Chapter 4).

46

CHAPTER 2

The Lactate Dehydrogenase-Elevating Virus Capsid Protein is a Nuclear-

Cytoplasmic Protein

Hakimeh Mohammadi1, Shayan Sharif

1, Raymond R. Rowland

2, Dongwan Yoo

3,*

1Department of Pathobiology, University of Guelph, Guelph, Ontario N1G 2W1, Canada.

2Department of Diagnostic Medicine and Pathobiology, Kansas State University,

Manhattan, KS 66506, USA.

3Department of Pathobiology, University of Illinois at Urbana-Champaign, 2001 South

Lincoln Avenue, Urbana, IL 61802, USA.

Published in: Archives of Virology. 2009.154(7):1071-80.

47

ABSTRACT

Arteriviruses replicate in the cytoplasm and do not require the nucleus function of

cell for virus multiplication in vitro. However, nucleocapsid (N) protein of two

arteriviruses, porcine reproductive respiratory syndrome virus (PRRSV) and equine

arteritis virus (EAV), has been observed to localize in the nucleus and nucleolus of virus-

infected and N gene-transfected cells, in addition to their normal cytoplasmic

distribution. In the present study, the N protein of lactate dehydrogenase-elevating virus

(LDV) of mice was examined for nuclear localization. Subcellular localization of LDV-N

was determined by tagging N with enhanced green fluorescence protein (EGFP) on the

N- and C-terminus. Both N-EGFP and EGFP-N fusion proteins localized to the nucleus

and nucleolus of gene-transfected cells. Labeled N also accumulated in the perinuclear

region, the site of virus replication. LDV-N sequence contains a putative ‘pat4’ type

nuclear localization signal (NLS) consisting of KKKK. To determine its functional

significance, a series of deletion constructs of N were generated and individually

expressed in cells. The results showed that the ‘pat4’ NLS was essential for nuclear

translocation. In addition, the LDV-N interacted with the importin-α and -β proteins,

suggesting that the LDV-N nuclear localization occurs via the importin-mediated nuclear

transport pathway. These results provide further evidence for the nuclear localization of

N as a common feature within the arteriviruses.

48

INTRODUCTION

Lactate dehydrogenase-elevating virus (LDV) is a murine virus causing lifelong

persistent viremia in mice with no apparent clinical signs except an increased level of the

plasma enzyme lactate dehydrogenase (Plagemann et al., 1995). LDV shows a limited in

vitro replication which is restricted to only specific murine primary macrophages and

presents a low level of virus production in culture (Brinton-Darnell et al., 1975; Brinton-

Darnell and Plagemann 1975; Stueckemann et al., 1982; Onyekaba et al., 1989a). LDV is

an RNA virus of 62-80 nm in diameter which belongs to the family Arteriviridae. This

family of viruses contains a single-stranded positive-sense RNA as the genome and

includes three other member viruses: porcine reproductive and respiratory syndrome

virus (PRRSV), equine arteritis virus (EAV), and simian hemorrhagic fever virus (SHFV)

(Snijder et al, 1998). The family Arteriviridae is grouped in the order Nidovirales along

with the Coronaviridae family. Arteriviruses infect macrophages in the host animal and

are responsible for a number of important diseases such as abortions and respiratory

disorders in pigs by PRRSV, persistent viremia in mice by LDV, abortion and arteritis in

horses by EAV, and hemorrhagic fever in monkeys by SHFV (Snijder and Spann, 2007).

The LDV genome is 14.1 kb in length dominated by two large open reading

frames ORF1a and ORF1ab, in the 5’ three-fourths of the genome and code for two large

polyproteins PP1a and PP1ab. The polyproteins are predicted to be proteolytically

cleaved into 12 cleavage products designated Nsp1 (non-structural protein 1) through

Nsp12. Downstream from ORF1ab, are ORF2 through ORF7, which encode at least 7

structural proteins; GP2a, GP2b (E), GP3, GP4, GP5, matrix (M) and nucleocapsid (N)

49

proteins (Snijder and Spann, 2007). The nucleocapsid protein of LDV is 115 amino acids

in length and is the most abundant protein within the virion.

Nidoviruses replicate exclusively in the cytoplasm. However, the N protein of

PRRSV, EAV and the coronaviruses, avian infectious bronchitis virus (IBV) and

transmissible gastroenteritis virus (TGEV), localize to the nucleolus (Rowland et al.,

1999; Tijms et al., 2002, Wurm et al, 2001). An interesting exception is the N protein of

SARS-CoV. One study showed that SARS-CoV N protein is localized exclusively in the

cytoplasm (Rowland et al, 2005), while another study has reported the SARS-CoV N

protein is present in the both cytoplasm and the nucleus/nucleolus (You et al, 2007).

These findings suggest that localization of N to the nucleolus may not be a general

property for all nidovirus N proteins. Once in the nucleus, the PRRSV and IBV N

proteins interact with fibrillarin and nucleolin, where they may be involved in the cell

cycle regulation (Yoo et al., 2003; Chen et al., 2002; Wurm et al., 2001). For nuclear

proteins, trafficking between cytoplasm and the nucleus is through the nuclear pore

complexes (NPC). While small molecules enter the nucleus by diffusion through NPC,

large molecules enter the nucleus by an energy-dependant pathway using a nuclear

localization signal (NLS) (Golrich and Kutay, 1999). The NLS on the cargo protein

interacts with importin proteins in the cytoplasm and then are shuttled across the NPC

and into the nucleus. Three types of NLSs have been identified. A “pat4” type NLS is a

stretch of four basic amino acids such as ‘KKKK’. A second type is “pat7” which is a

string of amino acids starting with proline (P) followed by any other two amino acids

which is followed by four amino acids of which at least three are basic residues. The third

type is a bipartite form. The bipartite NLS has two areas of basic residues separated by a

50

10-12 amino acids. The first group of basic residues includes two basic amino acids and

the second group contains 5 amino acids consisting of at least three basic residues (Nakai

and Kanehisa, 1992; Rowland and Yoo, 2003). In this report, we examined the N protein

nuclear localization of LDV and identified a functional ‘pat-4’ signal sequence essential

for N protein translocation into the nucleus.

MATERIALS AND METHODS

Cells, viruses, and bacterial strains: MARC-145 (Kim et al, 1993), a derivative of MA-

104 cells, HeLa cells, and NIH-3T3 cells were used for this study. Cells were maintained

in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% heat

inactivated fetal bovine serum (FBS, Invitrogen, Carlsbad, CA) in a humidified incubator

with 5% CO2 at 37ºC. PRRSV strain PA8 (Wootton et al., 2000) was grown in MARC-

145 cells. E. coli strains DH5α and XL-1 Blue (Stratagene, La Jolla, CA) were used for

gene cloning and mutagenesis, and strain BL-21 (Stratagene) was used for protein

expression.

Plasmid constructions: The LDV-N gene was amplified by PCR from a plasmid

containing the LDV genomic fragment, and using the BamHI sequence at both ends of

the primers, cloned into pEGFP-N1 and pEGFP-C1 (Clontech, Mountain View, CA)

upstream and downstream of the enhanced green fluorescence protein (EGFP) gene,

respectively. LDV-N was also subcloned using BamHI in pCITE-2C (Novagen) for in

vitro transcription and translation.

Mutant construction for LDV-N: Deletion constructs of LDV-N were made by PCR

using respective primer sets (Table 2.1) under the following conditions: preincubation for

51

Table 2.1. List of primers used for EGFP fusion constructions for LDV N.

Construct Primer pair Sequence a

LDV N-EGFP N-EGFP-FWD

N-EGFP-REV

5’-ATggatccATGTCTCAAAATAAGAAGAA-3’

5’-ATggatccGAAGCAGAAGAATTAGCAGAAG-3’

EGFP-LDV N EGFP-N-FWD

EGFP-N-REV

5’-ATggatccATGTCTCAAAATAAGAAGAA-3’

5’-ATggatccTTAAGCAGAAGAATTAGCAG-3’

LDV N- ΔC40 N-ΔC40-FWD

N- ΔC40-REV

5’-ATggatccATGTCTCAAAATAAGAAGAA-3’

5’-ATggatccGACAGCTTGGGCTGCTTCTTCTT-3’

LDV N- ΔC65 N-ΔC65-FWD

N- ΔC65-REV

5’-ATggatccATGTCTCAAAATAAGAAGAA-3’

5’-ATggatccGACGCCATAGGGAAGTGCAGCTT-3’

LDV N- ΔC70 N-ΔC70-FWD

N- ΔC70-REV

5’-ATggatccATGTCTCAAAATAAGAAGAA-3’

5’-ATggatccGAGAACAAGGTTACCAATGAGGC-3’

LDV N- ΔC82 N-ΔC82-FWD

N- ΔC82-REV

5’-ATggatccATGTCTCAAAATAAGAAGAA-3’

5’-ATggatccGATTTATTCTGTCCGGCATTGCG-3’

a Primers were designed based on sequence information available for LDV.

b Lowercase letters indicate BamHI recognition sequence.

52

5 min at 37ºC, denaturation for 30 sec at 95ºC, annealing for 1 min at 56ºC, and extension

for 1 min at 72ºC per cycle for 35 cycles, and final extension for 10 min at 72ºC in the

GeneAmp 2400 thermocycler (Perkin Elmer). PCR products were digested with BamHI

and subcloned into pEGFP-N1. Each construct was verified by restriction digestion and

nucleotide sequencing of the junction between N and EGFP. Cloning and DNA

manipulations were conducted according to the standard protocols (Sambrook and

Russell, 2001). For mutagenesis, PCR-based site-directed mutagenesis was performed by

overlapping extension using mismatch primers as previously described (Table 2.2)

(Wootton et al., 2001). PCR was carried out using 15 ng of pLDV-N-EGFP plasmid

DNA, 300 ng of each forward and reverse primer, 1 mM dNTPs, 1X buffer [10 mM KCl,

10 mM (NH4)2SO4, 20 mM Tris-HCl (pH 8.8), 2 mM MgSO4, 0.1% Triton X-100], and

2.5 units of Pfu DNA polymerase (Stratagene) for 16 cycles as follows: denaturation at

94°C for 30 sec, primer annealing at 56°C for 1 min, and primer extension at 68°C for 8

min. The PCR products were digested with DpnI and transformed to E. coli XL1-Blue.

Transformants were randomly selected for plasmid DNA preparation using a QIAprep

spin miniprep kit (QIAGEN, Santa Clarita, CA), and mutagenized sequences were

verified by nucleotide sequencing in both directions.

Fluorescent microscopy: Expression of N-GFP fusion proteins was determined in HeLa

cells by transfection using Lipofectin (Invitrogen) according to the manufacturer’s

instruction. Cells were grown on microscope coverslips and transfected with GFP fusion

constructs of LDV-N. PRRSV N gene and PRRSV N-GFP fusion constructs were

included as a positive control. At 24 hrs post-transfection, cells were fixed with methanol,

blocked with 1% bovine serum albumin (BSA) in PBS (phosphate buffered saline) for 30

53

min, and visualized for fluorescence using a fluorescence microscope (model AX70,

Olympus). For PRRSV-infected cells, N protein was stained with MAb SDOW-17

(Nelson et al., 1993) at a dilution at 1:400 in 1% BSA-PBS, followed by goat anti-mouse

antibody conjugated with AlexaFluor 488 (Molecular Probes) in dilution of 1:100. After

5 washes with PBS, cells were stained with DAPI (4',6-Diamidino-2-phenylindole

dihydrochloride, Sigma) for 5 min, washed again, mounted on a slide using 40 μl of the

mounting buffer (60% glycerol, 0.1% sodium azide in PBS), and visualized for

fluorescence.

GST-fusion protein expression and coupling with Sepharose beads: Mouse importin-

α and importin-β genes were cloned in pGEX-3X (Amersham, Piscataway, NJ) and

expressed as GST (glutathione-S-transferase)-importin-α and GST-importin-β fusion

proteins. One ml of Luria-Bertani medium containing 100 g/ml of ampicillin was

inoculated with 1/100 of an overnight culture. When optical density of the culture reaches

0.6 at 600 nm, IPTG (isopropyl-β-D-thiogalactopyranoside) was added to the final

concentration of 1 mM and the culture was further incubated for 3 h. Bacterial cells were

collected by centrifugation at 6,000 RPM for 10 min 4oC (Avanti J30I, Beckman) and

resuspended in 10 ml of PBS. The bacterial suspension was sonicated on ice three times

(model W-385; Ultrasonics Inc.), each time for 30 sec with 2 sec intervals. The

suspension was incubated with 1% Triton X-100 for 30 min at 4oC with occasional

agitation and centrifuged at 10,000 RPM for 10 min at 4oC to remove the insoluble

fractions and cell debris. The supernatant which contained expressed GST-fusion proteins

was incubated with 100 l of glutathion-Sepharose 4B beads (Amersham) in 50% slurry

overnight at 4oC with constant agitation. GST-fusion proteins bond to beads were

54

centrifuged and washed 5 times in PBS containing 1% Triton X-100. The collected beads

were then resuspended in a final volume of 250 l of incubation buffer (20 mM Tris-HCl

[pH 7.5], 100 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 5 mM dithiothreitol, 0.5% NP-40, 1

mM PMSF, 5% glycerol). This resulted in 20% slurry of beads to use for GST pull-down

assay. For production of radiolabelled N protein, the LDV-N gene was cloned into pCITE

(Novogen) and translated using TNT® Quick Coupled in vitro Transcription/Translation

system (Promega, Madison, WI) in the presence of [35

S]methionine (EasyTag EXPRESS

protein-labeling mix of [35

S]methionine and [35

S]cysteine; specific activity 407 MBq/ml,

Perkin-Elmer) according to the manufacturer’s instruction. Based on the quantity as

measured by SDS-PAGE, approximately equal amounts of Sepharose-coupled GST

fusion proteins and LDV-N were incubated overnight at 4°C with constant agitation. The

Sepharose beads were washed five times with PBS, boiled for 5 min in SDS sample

buffer (10 mM Tris-HCl [pH 6.8], 25% glycerol, 10% SDS, 10% β-mercaptoethanol, and

0.12% [wt/vol] bromophenol blue), and analyzed by 12% SDS-PAGE. Gels were dried

and exposed for image analysis by Phosphorimager (Molecular Dynamics).

RESULTS

Subcellular localization of EGFP-tagged LDV-N: The LDV nucleocapsid protein was

expressed as an EGFP-N fusion protein in HeLa cells and examined for subcellular

distribution. Cells were counterstained with DAPI for nuclear staining. EGFP alone was

distributed throughout transfected cells (Fig. 2.1B). For the purpose of comparisons,

localization of the PRRSV N protein in virus-infected MARC-145 cells is shown (Fig.

2.1C). Detection of PRRSV N by N-specific MAb SDOW17 showed the accumulation of

55

56

Fig. 2. 1. Subcellular localization of LDV-N protein. HeLa cells grown on the coverslips

were transfected with EGFP-LDV-N fusion protein genes using Lipofectin. At 24 hrs

post-transfection, cells were fixed with methanol and stained with DAPI for nuclear

staining. PRRSV-infected MARC-145 cells were stained with SDOW-17 MAb, followed

by staining with goat anti-mouse Ab conjugated with AlexaFluor 488 and finally were

stained with DAPI for nuclear staining. Cells on the coverslips were visualized by

fluorescence microscopy (model AX70, Olympus). Panels: A, HeLa cells without

transfection; B, HeLa cells with the pEGFP-N1 empty vector; C, PRRSV-infected

MARC-145 cells stained with the N-specific MAb SDOW-17; D, PRRSV-infected

MARC-145 cells stained with bovine coronavirus spike protein-specific MAb; E, HeLa

cells with PRRSV-N gene stained with SDOW-17; F, HeLa cells with PRRSV-N-GFP;

G, HeLa cells with LDV-N-EGFP; H, HeLa cells with EGFP-LDV-N.

57

N in the nucleus and nucleolus in addition to the normal cytoplasmic distribution. In

PRRSV N and PRRSV N-GFP transfected cells, N accumulated in the nucleus (Fig. 2.1E

and 2.1F), and similarly, both LDV-N-EGFP and LDV-EGFP-N fusion proteins showed

the accumulation of fluorescence in the nucleus (Fig. 2.1G and 2.1H), with a distribution

pattern similar to PRRSV N, indicating that LDV-N localized both in the cytoplasm and

the nucleolus.

Identification and mapping of NLS in the LDV-N protein: The sequence analysis of

the LDV-N protein identified the presence of a single basic region of KKK which

resembled a ‘pat-4’ type NLS located between amino acid positions 38-41 (Fig. 2.2A).

The location of this lysine-rich region corresponded to the ‘pat-7’ motif of PRRSV N

protein. To determine the functional significance of the ‘pat-4’ motif of LDV-N, a series

of deletion mutants were made by progressively deleting from the C-terminus of N (Fig.

2.2B). Mutant constructs were then transfected into cells and their fluorescence was

examined. LDV-N-C40-EGFP, LDV-N-C65-EGFP, and LDV-N-C70-EGFP showed the

fluorescence in the nucleus and nucleolus (Fig. 2.2(C); panels B, C, D).

In contrast, LDV-N-C82-EGFP in which the putative motif was deleted showed the

presence of cytoplasmic fluorescence but the absence of nuclear fluorescence (panels E).

These results indicate that the predicted ‘pat-4’ motif was indeed a functional NLS that

translocates LDV-N to the nucleus and nucleolus in cells.

To further confirm the function of predicted NLS and to identify core amino acids

essential for this function, individual residues of ’38-KKKK-41’ were substituted with

glycine (G) at single or multiple sites (Table 2.2), and the individual mutants were then

expressed in HeLa cells (Fig. 2.4A). This study showed that two ‘K’s at positions 38 and

58

59

Fig. 2. 2. Identification of an NLS motif in the LDV-N protein and its functional

mapping. A, a predicted ‘pat-4’ type nuclear localization signal in the LDV-N protein

identified by the PSORT II program. The PRRSV N protein sequence is aligned for

comparison with LDV-N. Dashes indicate deletions. Bold letters with underlines indicate

‘pat-4’ (for LDV) and ‘pat-7’ (for PRRSV) type NLS sequences with their amino acids

positions. B, schematic presentation of LDV-N deletion mutants fused with EGFP.

Darkened areas represent predicted NLS at amino acid positions 38-41. C, subcellular

localization of LDV-N mutants, upper and lower images are GFP expression of the LDV

N constructs and the overlay of GFP and DAPI staining of the same area respectively.

Panels (A), wild type LDV-N-EGFP; (B), deletion constructs LDV-N-C40-EGFP; C,

LDV-N-C65-EGFP; D, LDV-N-C70-EGFP; E, LDV-N-C82-EGFP.

60

Table 2.2. Site-directed mutagenesis of LDV N NLS motif and subcellular localization of

NLS mutants.

Construct Sequence of NLSa

Localizationb

Cy Nu No

LDV-N 38-KKKK-41 ++++ ++ ++++

LDVN- K38G 38-GKKK-41 ++++ ++++ +++

LDV-N- K39G 38-KGKK-41 ++++ +++ +

LDV-N- K40G 38-KKGK-41 ++++ ++ +

LDV-N- K41G 38-KKKG-41 ++++ ++++ +++

LDV-N- K38,39G 38-GGKK-41 ++++ ++++ +

LDV-N- K38,40G 38-GKGK-41 ++++ ++ +

LDV-N- K38,41K 38-GKKG-41 ++++ +++ +

LDV-N- K39,40G 38-KGGK-41 ++++ +++ ++

LDV-N- K39,41G 38-KGKG-41 ++++ +++ +

LDV-N- K40,41G 38-KKGG-41 ++++ +++ +/-

LDV-N- K39,40,41G 38-KGGG-41 ++++ +++ +/-

LDV-N- K38,39,40G 38-GGGK-41 ++++ +++ +/-

LDV-N- K38,39,40,41G 38-GGGG-41 ++++ - -

aIndividual lysines (underlined) in the NLS were substituted with glycines.

bFor scoring of the fluorescence signal for each mutant construct, 100 transfected cells

were counted presented as + or -. Each one ‘+’ represents fluorescence of 25 transfected

cells either in the cytoplasm (Cy), nucleus (Nu), or nucleolus (No).

61

Fig. 2. 3. Subcellular localization of NLS mutants. Panels A through D, single

substitutions of LDV-N (LDV-N-K38G, LDV-N-K39G , LDV-N-K40G, and LDV-N-

K41G, respectively) panels E through J are double mutations in NLS (LDV-N-K38,39G ,

LDV-N-K38,40G, LDV-N-K38,41K , LDV-N-K39,40G , LDV-N-K39,41G , LDV-N-

K40,41G, respectively). Panels K and L are triple mutations in NLS (LDV-N-

K39,40,41G and LDV-N-K38,39,40G). (A), GFP staining; (B), the same cells in (A)

overlaid with DAPI staining.

62

Fig. 2. 4. Subcellular localization of the NLS-null LDV-N-EGFP mutants in HeLa cells

(A) and NIH-3T3 murine cells (B). The NLS (38-KKKK) of LDV-N was mutated to 38-

GGGG-41 to knock out the NLS function. The NLS-null LDV-N gene was transfected to

HeLa cells or NIH-3T3 cells followed by staining with DAPI for fluorescent microscopy.

LDV-N-wt, wild type LDV-N-EGFP; LDV-N-null, NLS-null mutant LDV-N-EGFP;

PRRSV-N, porcine reproductive respiratory syndrome virus wild type N protein.

63

40 function as core amino acids for nucleolar translocation of N and ‘K’s at positions 38,

40, and 41 play a crucial role for this function (Table 2.2 and Fig. 2.3). The mutant N in

which all four ‘K’s were substituted with ‘G’s was exclusively cytoplasmic and did not

localize to either nucleus or nucleolus (Fig. 2.4), confirming that the pat-4’ motif was

indeed the functional NLS for LDV-N protein for nuclear localization. Since LDV is a

murine virus, the function of NLS and the nuclear localization of LDV-N were confirmed

in mouse cells using the same constructs (Fig. 2.4B). NIH-3T3 cells were transfected

with either wild-type LDV-N or LDV-N NLS-null mutant construct and examined for

fluorescence. The wild-type LDV-N protein was clearly nuclear-localized, and the LDV-

N NLS-null mutant was cytoplasmic without distinct nuclear staining, indicating that the

LDV-N nuclear localization is not cell type-dependent but is rather the nature of the N

protein.

LDV-N interaction with importin proteins: To determine the basis for LDV-N nuclear

localization, interactions between N and importin proteins were determined by GST pull-

down assay. Importin-α and importin-β were individually expressed in E.coli as a GST-

fusion protein and coupled to glutathione-Sepharose beads. The radiolabelled LDV-N

protein was then synthesized in vitro by transcription translation, and importin-coupled

GST beads were incubated with the radiolabelled N protein. Unbond proteins were

washed off and the bond proteins were resolved by SDS-PAGE and autoradiography. As

with the PRRSV N protein which was previously shown to interact with both importin-α

and -β proteins (Rowland et al., 2003; Fig. 2.5, lanes 7 and 9), both importin-α and

importin-β specifically bond LDV-N (lanes 8 and 10), while GST alone did not bind to

both PRRSV-N and LDV-N proteins (lanes 5 and 6). This study demonstrates the specific

64

interaction of LDV-N with both importin proteins, and suggests that LDV-N nuclear

localization may be importin-mediated.

DISCUSSION

Nuclear localization of capsid protein has been studied for several RNA viruses

that restrict their replication in the cytoplasm, and capsid proteins of Semliki forest virus,

dengue fever virus, and hepatitis C virus have been shown to localize in the nucleus and

nucleolus (Favre et al., 1994; Yasui et al., 1998, Wang et al., 2002). Nidoviruses also

replicate exclusively in the cytoplasm of infected cells (Brayton et. al, 1981), but N

protein has been shown to localize in the nucleus and nucleolus for PRRSV, EAV, IBV,

MHV and TGEV (Hiscox et al, 2001; Wurm et al, 2001; Rowland et al., 1999; Tijms et

al., 2002). The N protein of PRRSV contains two putative NLS motifs, ‘pat 4’ and ‘pat

7’, and the “pat 7” located at positions 41-47 has been shown to be the active and primary

NLS for N (Rowland et al., 2003). In the present study, we showed that the LDV-N

protein localized to the nucleus and nucleolus in N-EGFP gene transfected cells, and

using mutant constructs showed that the nuclear localization of N was dependent on a

‘pat-4’ motif located at positions 38-41 (Fig. 2. 2A; 2C). The location of ‘pat-4’ motif in

LDV-N is similar to the region where the functional ‘pat 7’ motif is located in PRRSV N.

We further identified two lysine residues at positions 38 and 40 in LDV-N that form core

residues for nuclear transport of N. For PRRSV N, amino acids at position 43 and 44 are

critical residues for nuclear localization (Rowland et al., 2003; Lee et al., 2006a; Pei et

al., 2008). For a protein to localize in the nucleus, the cellular chaperone importin-α

recognizes the NLS on the cargo protein and subsequently, importin-β binds to the

65

Fig. 2. 5. Interactions of LDV-N with mouse importin proteins by GST-pull down assays.

Importin-α- and importin-β-glutathione-S-transferase (GST) fusion proteins were

expressed in E-coli and coupled to glutathion-Sepharose 4B beads. Beads-bound proteins

were incubated overnight at 4°C with in vitro-translated radiolabelled N protein. After

washing, bound proteins were visualized by 12% SDS-PAGE and autoradiography. Left

panel, in vitro translation of vector control (lane 2), PRRSV-N (lane 3), and LDV-N (lane

4). Middle and right panels, GST pull-down assays for LDV-N by importin-α (lane 8)

and importin-β (lane 10), respectively.

66

importin-α/cargo protein complex, transporting the complex in the presence of Ran-GDP

to NPC on the nuclear membrane. We found that LDV-N binds to importin-α (Fig. 2.5,

lane 8), which is consistent with the utilization of the classical pathway for nuclear

transport. It is interesting to note that LDV-N also binds to importin-β (Fig. 2.5, lane 10).

PRRSV N also binds to both importin-α and importin-β (Rowland et al, 2003) and

several other viral proteins have been reported to interact with both importin-α and -β.

The polyomavirus capsid protein requires the interaction with both importin-α and -β to

enter the nucleus (Qu et al., 2004), and SV40 VP3 also interacts with both importin

proteins for nuclear translocation (Nakanashi et al, 2002). The HIV Rev protein binds

both importin-α and -β, but its binding to importin-α is NLS-independent while the

importin- β binding is NLS-dependent (Henderson et al., 1997). For PRRSV, two

nucleolar proteins, nucleolin and B23, have been identified to bind to N (unpublished

data), and thus a possibility for the LDV-N nuclear transport is the incorporation of

importin-dependent pathway with a ‘piggy-backing’ mechanism, in which N binds to a

nuclear targeted cellular protein in addition to importin proteins.

Since N proteins of PRRSV, EAV, and LDV all localize to the nucleus and

nucleolus, the nuclear role of N during infection is of major interest. The core protein of

hepatitis C virus is believed to modify cellular function by preventing translocation of

host cell proteins to the nucleus (Isoyama et al., 2002; Suzuki et al., 2005), and the N

protein of TGEV has been suggested to play a role in the disruption of cell division

(Wurm et al., 2001). For PRRSV N, disruption of NLS does not affect virus replication in

cell culture, but in pigs, NLS-null viruses are attenuated in their virulence (Lee et al.,

2006a, 2006b; Pei et al., 2008). The NLS motif of PRRSV N was associated with strong

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selective pressure for reacquisition of the nuclear function in vivo, suggesting the

involvement of N nuclear localization in the viral pathogenesis. N proteins of several

member viruses in the order Nidovirales colocalize and interact with fibrillarin and

nucleolin in the nucleolus, supporting the hypothesis that N modulates the host cell

ribosomal biogenesis and cell cycle progression (Chen et al., 2002; Hiscox, 2007). In

summary, we show that LDV-N contains a functional ‘pat-4’ type NLS at positions 38-41

and localizes in the nucleolus of cell via the importin-mediated pathway. The capsid

protein nuclear localization may be a common feature in the Arteriviridae family, which

may play a common role for replication and pathogenesis of this group of viruses.

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CHAPTER 3

Nuclear Localization of Equine Arteritis Virus and Simian Hemorrhagic Fever Virus

Nucleocapsid Proteins

69

ABSTRACT

In the present study, the subcellular localization of nucleocapsid (N) proteins of

two member viruses in the Arteriviridae family, equine arteritis virus (EAV) and simian

hemorrhagic fever virus (SHFV) were examined by immunofluorescence microscopy.

EAV N and SHFV N proteins were fused with the enhanced green fluorescence protein

(EGFP) on the N- or C-terminus, and the cellular distribution of fusion proteins was

examined in gene-transfected cells. Both N-EGFP and EGFP-N fusion proteins of EAV

and SHFV showed accumulation of fluorescence in the nucleus and nucleolus of gene-

transfected cells. The EAV N gene contains a putative ‘bipartite’ type nuclear

localization signal (NLS) at position 4-20, and the SHFV N gene has a potential ‘pat-7’

type NLS at amino acid positions 22-28. To determine the functional significance of the

NLS-like sequences, a series of deletion mutants were made by progressively deleting

amino acids from the C- and N-termini of both EAV N and SHFV N proteins and the

mutant proteins were individually expressed in HeLa cells. The results show that the

‘pat7’ NLS was essential for nuclear translocation of SHFV N protein, while EAV N

protein required the entire motif containing 4-20 N-terminal residues for nuclear

localization. Furthermore, the interaction of SHFV N protein with importin-α indicates

the use of the importin-mediated nuclear transport pathway. EAV N protein interacted

with both importin-α and importin-β proteins, which indicates that the importin mediated

pathway is likely involved in its nuclear transfer. Taken together, this study indicates that

nucleolar localization of the N protein is a common feature of the Arteriviridae family

and possibly plays a significant role in the virus life cycle of this family.

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INTRODUCTION

Arteriviruses include four members of enveloped RNA viruses containing a

single-stranded positive-sense RNA genome of 12.7 to 15.7 kilobases (kb) in size with

the 5` cap structure and 3` polyadenylated tail (Snijder and Swaan, 2007). Based on the

similarity of the mechanism for their genome replication, the Arteriviridae family has

been placed along with the Coronaviridae family, into the order Nidovirales (Snijder and

Spann, 2007; Gorbalenya et al, 2006).

Equine arteritis virus (EAV) and simian hemorrhagic fever virus (SHFV) are two

member viruses of the Arteriviridae family and cause viral arteritis in horses and

hemorrhagic fever in monkeys, respectively. EAV causes respiratory and reproductive

failures in horses, donkeys, and mules resulting in economic losses for the equine

industry in North America (Timoney and McCollum, 1993). The causative agent for viral

arteritis is EAV which was first identified in 1957 in Bucyrus, Ohio from lungs of sick

horses affected by this disease (Doll et al, 1957; Bryans et al 1957). SHFV causes acute,

febrile and fatal disease with a mortality rate of up to 100% in rhesus monkeys (Palmer et

al, 1968). It was first isolated from a fatal hemorrhagic fever of macaque monkeys in

1968 (Madden et al., 1978, Trousdale et. al 1975).

EAV is the prototype virus of the Arteriviridae family (Snijder and Meulenberg,

1998). Its genome is an infectious single-stranded RNA 12.7 kb in size which contains 9

open reading frames (ORF). ORF1a and ORF1ab are translated directly from the viral

genome, and the two polyproteins are cleaved to produce 18 non-structural proteins.

These proteins are involved in viral genome transcription and replication. ORFs 2a, 2b, 3,

4, 5, 6, and 7 code for structural proteins E, GP2 (GP2b/Gs), GP3, GP4, GP5 (GL), M, and N

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respectively. The M and GP4 are two major envelope proteins, and E, GP2b, GP3, and

GP4 are the minor envelop proteins (Snijder and Spann, 2007; de Vries et al, 1992;

Wieringa et al, 2004; Molenkamp et al, 2000b). N protein is the most abundant protein of

EAV involved in virion RNA encapsidation, while being dispensable for viral genome

replication (de Vries et al, 1992; Wieringa et al., 2004; Molenkamp et al, 2000b). EAV N

protein is a phosphoprotein of 110 amino acids with the molecular weights of 12-14 kD

(Zeeger et al, 1976; de Vries et al, 1992).

The genome of SHFV is also an infectious RNA of 15.7 kb in size. It contains two

non-structural and 9 structural ORFs. Like other members of arteriviruses, non-structural

proteins of SHFV are also translated from the two largest ORFs; ORF1a and ORF1b.

ORFs 2a, 2b, 3, 4a, 4b, 5, 6, 7, 8, and 9 code for structural proteins Gp2a, Gp2b, Gp3, E,

Gp4, Gp5, Gp6, Gp7(p54), M, and N, respectively. ORF9 encodes nucleocapsid (N)

protein which is one of the most abundant structural proteins of SHFV with 115 amino

acids in length (Sagripanti 1984).

Previous studies have shown that the N protein of PRRSV localizes to the

cytoplasm and the nucleus/nucleolus of PRRSV-infected and N-gene transfected cells

(Rowland et al., 1999). EAV N also seems to localize in the nucleus and nucleolus of

infected cells (Tijms et al., 2002). A similar observation has been reported for the N

protein of lactate dehydrogenase-elevating virus (LDV) in human and mouse cells

expressing LDV N (Mohammadi et al., 2009). Similar cellular distribution of the N

proteins have been reported for coronaviruses infectious bronchitis virus

(IBV),

transmissible gastroenteritis virus (TGEV), and severe acute respiratory syndrome

(SARS) virus (Wurm et al, 2001, You et al, 2007). As in Coronaviridae, viruses in the

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Arteriviridae family replicate in the cytoplasm of infected cells and do not need the

nuclear function of infected cells for their life cycle. The finding of the nucleolar

localization of N protein in some arteriviruses including PRRSV and EAV suggests that

the nucleolar localization of N protein may be a common feature in nidoviruses with an

important function for viral replication in addition to its structural role for virion

assembly of these viruses. Despite the investigation using PRRSV and LDV, the cellular

distribution of SHFV N and EAV N has yet to be investigated.

The objectives of this study were to determine the SHFV N transport to the

nucleus, to define the nuclear localization signal (NLS) for EAV and SHFV N proteins,

and to elucidate the mechanism of their transport to the nucleus. This study shows that

the nuclear localization of N is a conserved feature of arteriviruses and indicates that the

nucleolar localization of the N protein may play an important role in the pathogenesis of

diseases caused by these viruses.

MATERIALS AND METHODS

Cells, viruses, and bacterial strains. Baby hamster kidney cells (BHK-21), MARC-145

(Kim et al, 1993) which is a derivative of the MA104 cell line, COS-1, and HeLa cells

were used in this study. Cells were maintained in Dulbecco’s modified Eagle’s medium

(DMEM) supplemented with 10% heat inactivated fetal bovine serum (FBS; GIBCO

BRL) in a humidified incubator with 5% CO2 at 37ºC. EAV strain Bucyrus (Doll et al.,

1957) and PRRSV strain PA8 (Wootton et al., 2000) were grown in MARC-145 and

BHK-21 cells, respectively. Escherichia coli (E. coli) strains XL-1 Blue (Stratagene, La

73

Jolla, CA), and DH5α were used for gene cloning and mutagenesis, and E. coli strain BL-

21 (Novagen) was used for protein expression.

Antibodies. A mouse MAb SDOW-17 (IgG isotype) against the PRRSV N (Nelson et al.,

1993) was used to stain for PRRSV N protein. A mouse MAb 3E2 (IgG type) against

EAV N was a generous gift from Dr. Udeni Balasuriya (Gluck Equine Research Center,

University of Kentucky, Lexington, USA) and used to stain for EAV N protein. Goat

anti-mouse antibody conjugated with AlexaFluor 488 was purchased from Molecular

Probes and used as a secondary antibody.

Plasmid constructions. The EAV N gene was amplified from pEAV030 containing the

full genome sequence of EAV (van Dinten et al., 1997) by PCR using the specific

primers containing a BamHI recognition sequence at the 5’ end (Table 3.1). The PCR

product was subcloned in-frame into pEGFP-N1 and pEGFP-C1 (Clontech Inc.)

upstream and downstream of the enhanced green fluorescence protein (EGFP) gene,

respectively. In order to clone the EAV N gene upstream of EGFP, the reverse primer

was designed to delete the stop codon of EAV N and the PCR product was subcloned in

frame with EGFP. SHFV N gene was PCR-amplified from a cDNA library of SHFV

genome (provided by Dr. E. Snijder, Leiden University Medical Center, Leiden,

Netherlands) using specific primers (Table 3.2) and sub-cloned upstream and

downstream of EGFP in the pEGFP-N1 and pEGFP-C1 in-frame, respectively. In order

to construct the SHFV N-EGFP in frame, the reverse primer of N-EGFP was designed to

eliminate stop codon of the SHFV N gene. Both EAV N and SHFV N were subcloned

into BamHI site of pCITE-2C (Novogen) for in vitro transcription and translation.

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Table 3.1. List of primers used for EGFP fusion constructions for EAV N.

Construct Primer pair Sequence a

EAV N-EGFP N-EGFP-FWD

N-EGFP-REV

5’-ATggatccATGGCGTCAAGACGATCACGT-3’

5’-ATggatccGACGGCCCTGCTGGAGGCGCAA-3’

EGFP-EAV N EGFP-N-FWD

EGFP-N-REV

5’-ATggatccATGGCGTCAAGACGATCACGT-3’

5’-ATggatccTTACGGCCCTGCTGGAGGCGCA-3’

EAV N- ΔC40 N-ΔC40-FWD

N- ΔC40-REV

5’-ATggatccATGGCGTCAAGACGATCACGT-3’

5’-GTggatccGATTGTACGTTCGACGAAAGGGT-3’

EAV N- ΔC70 N-ΔC70-FWD

N- ΔC70-REV

5’-ATggatccATGGCGTCAAGACGATCACGT-3’

5’-GTggatccGAGGGCGGTTTGCGGACCCGCAT-3’

EAV N- ΔC87 N-ΔC87-FWD

N- ΔC87-REV

5’-ATggatccATGGCGTCAAGACGATCACGT-3’

5’-TAggatccGAGCTTGTAGGCTGTCGCCGC-3’

EAV N- ΔN20 N-ΔN20-FWD

N- ΔN20-REV

5’-CGggatccATGCCTACAAGCTACAATGACCT-3’

5’-ATggatccGACGGCCCTGCTGGAGGCGCAA-3’

a Primers were designed based on sequence information available for EAV.

b Lowercase letters indicate BamHI recognition sequence.

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Table 3.2. List of primers used for EGFP fusion constructions for SHFV N.

Construct Primer pair Sequence a

SHFV N-EGFP N-EGFP-FWD

N-EGFP-REV

5’-GGTggatccATGGCTGGCAAACCAAAAAC-3’

5’-GTggatccGAGGTGGAGGAGGTGACCTT-3’

EGFP-SHFV N EGFP-N-FWD

EGFP-N-REV

5’-GGTggatccATGGCTGGCAAACCAAAAAC-3’

5’-AGTggatccCTAGGTGGAGGAGGTGACCTTC-3’

SHFV N- ΔC20 N-ΔC20-FWD

N- ΔC20-REV

5’-CGggatccATGGCTGGCAAACCAAAA-3’

5’-GTggatccGAAGCAAAATTGATTCT-3’

SHFV N- ΔC40 N-ΔC40-FWD

N- ΔC40-REV

5’-CGggatccATGGCTGGCAAACCAAAA-3’

5’-GTggatccGAGATCAGCAGCTGTTT-3’

SHFV N- ΔC70 N-ΔC70-FWD

N- ΔC70-REV

5’-CGggatccATGGCTGGCAAACCAAAA-3’

5’-GTggatccGATAGAGGCTTGTGGAC-3’

SHFV N- ΔN30 N-ΔN30-FWD

N- ΔN30-REV

5’-CGggatccATGCGTAGAGCTGCTCCT-3’

5’-GTggatccGAGGTGGAGGAGGTGACCTT-3’

a Primers were designed based on sequence information available for SHFV.

b Lowercase letters indicate BamHI recognition sequence.

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Deletion construction and site-directed mutagenesis. Deletion constructs of EAV N

and SHFV N were made by PCR using respective primer sets (Tables 3.1 and 3.2) under

the following amplification conditions: preincubation for 5 min at 37ºC followed by 30

sec at 95ºC for denaturation, 1 min at 56ºC for annealing, and 1 min at 72ºC for extension

for 35 cycles, and the final extension for 10 min at 72ºC (GeneAmp 2400 thermocycler,

Perkin Elmer). Individual PCR products were digested with BamHI and subcloned into

the BamHI site of pEGFP-N1. Each construct was verified by restriction digestion and

nucleotide sequencing of the junction between N and EGFP. Cloning and DNA

manipulations were conducted according to the standard protocols (Sambrook and

Russell, 2001). To introduce mutations to NLS, PCR-based site-directed mutagenesis was

performed by overlapping extension using primer sequences for each mutation (Wootton

et al., 2001). PCR was performed using 15 ng of pEAV N-EGFP and pSHFV N-EGFP

plasmid DNA, 300 ng of forward and reverse primers (Tables 3.2 and 3.4), 1 mM dNTPs,

1X buffer [10 mM KCl, 10 mM (NH4)2SO4, 20 mM Tris-HCl (pH 8.8), 2 mM MgSO4,

0.1% Triton X-100], and 2.5 units of Pfu DNA polymerase (Stratagene) for 16 cycles

under the following parameters: denaturation at 94°C for 30 sec, primer annealing at

56°C for 1 min, and primer extension at 68°C for 8 min. Then, DpnI was added to PCR

products for 30 min and the digested DNA was transformed to E. coli XL1-Blue.

Colonies were randomly selected for plasmid DNA preparation using a QIAprep spin

miniprep kit (QIAGEN, Santa Clarita, Calif.) and sequencing. Nucleotide sequencing

was done in both directions to verify introduced mutations.

Fluorescence microscopy. Expression of N-GFP fusion proteins was determined in

HeLa cells by transfection using Lipofectin (Invitrogen) according to the manufacturer’s

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instruction. For SHFV N-GFP fusion constructs COS-1 and HeLa cells were used for

transfection. Cells were grown on microscope coverslips and transfected with EAV N-

GFP and SHFV N-GFP fusion constructs using Lipofectin according to the

manufacturer’s instructions. PRRSV N gene and PRRSV N-GFP fusion constructs were

included as a positive control. BHK-21 cells were infected with EAV at a MOI of 1. One

group of cells was treated with 10 µM of leptomycin-B (LMB). At 24 h post-transfection

and at 2, 4, 6, 8, 10, 12, 24, 36, and 48 h post-infection, cells were examined for EAV N

protein distribution. Briefly, cells were fixed with methanol, blocked with 1% bovine

serum albumin (BSA) in PBS (phosphate buffered saline) for 30 min, stained with DAPI

(4', 6-Diamidino-2-phenylindole dihydrochloride, Sigma) for 5 min, washed again,

mounted on a slide using 40 μl of the mounting buffer (60% glycerol, 0.1% sodium azide

in PBS), and visualized for GFP fluorescence by fluorescent microscope (model AX70,

Olympus). For PRRSV-infected and EAV-infected cells, they were stained with MAb

SDOW-17 (Nelson et al., 1993) and MAb 3E2 at a dilution at 1:400 in 1% BSA-PBS,

followed by goat anti-mouse antibody conjugated with AlexaFluor 488 (Molecular

Probes) at the dilution of 1:100. After 5 times washing with PBS, cells were stained with

DAPI for 5 min, washed again, mounted on a slide using 40 μl of the mounting buffer

and visualized for fluorescence by fluorescence microscope.

Protein expression in E. coli and coupling with Sepharose beads. Plasmids containing

the murine importin genes, mImp-α (PTAC58) and mImp-β (PTAC97), were used to

express the gutathione-S-transferase T-importin-α and -β fusion proteins (GST-Impα and

-β) in E. coli strain BL-21 (Rowland et al., 2003). One ml of Luria-Bertani medium

containing 100 g/ml of ampicillin was inoculated with 1/100 of an overnight culture.

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When the optical density of bacterial culture reached 0.6 at 600 nm, IPTG (isopropyl-β-

D-thiogalactopyranoside) was added to the final concentration of 1 mM and the culture

was further incubated for 3 hours. The bacterial cells were collected by centrifugation at

6,000 RPM for 10 min 4oC (Beckman Avanti J30I centrifuge) and resuspended in 10 ml

of PBS. The bacterial suspension was sonicated (model W-385; Ultrasonics Inc.) three

times, each time for 30 sec with 2 sec intervals. The suspension was incubated with 1%

Triton X-100 for 30 min at 4oC with occasional agitation and centrifuged at 10,000 RPM

for 10 min at 4oC to remove the insoluble fractions and cell debris. The supernatant

which contained expressed GST-fusion proteins was incubated with 100 l of glutathion-

Sepharose 4B beads (Pharmacia) in 50% slurry overnight at 4oC with constant agitation.

GST-fusion proteins bound to beads were centrifuged and washed 5 times in PBS

containing 1% Triton X-100. The collected beads were then resuspended in a final

volume of 250 l of incubation buffer (20 mM Tris-HCl [pH 7.5], 100 mM KCl, 2 mM

CaCl2, 2 mM MgCl2, 5 mM dithiothreitol, 0.5% NP-40, 1 mM PMSF, 5% glycerol). This

resulted in 20% slurry of beads to use for GST pull-down assay.

GST pull-down assay. To produce radiolabeled EAV N and SHFV N proteins, each N

gene was cloned into pCITE (Novogen) and translated using TNT® Quick Coupled in

vitro Transcription/Translation system (Promega) in the presence of 50 μCi/ml

[35

S]methionine (EasyTag EXPRESS protein-labeling mix of [35

S]methionine and

[35

S]cysteine; specific activity 407 MBq/ml, Perkin-Elmer) according to the

manufacturer’s instruction. The quantity of GST fusion proteins and translated EAV N

and SHFV N was measured by SDS-PAGE. Equal amounts of radiolabeled EAV N and

SHFV proteins were incubated with the GST fusion proteins bound to glutathione-

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Sepharose beads in 20% slurry in binding buffer separately (20 mM Tris-HCl [pH 7.5],

100mM KCl, 2 mM CaCl2, 2 mM MgCl2, 5 mM dithiothreitol, 0.5% NP-40, 1 mM

PMSF, 5% glycerol) to the final volume of 400 μl overnight at 4°C with constant

agitation. The beads were washed five times with PBS, boiled for 5 min in SDS sample

buffer (10 mM Tris-HCl [pH 6.8], 25% glycerol, 10% SDS, 10% β-mercaptoethanol, and

0.12% [wt/vol] bromophenol blue), and analyzed by 12% SDS-PAGE. Gels were dried

and exposed to Phosphorimager (Molecular Dynamics) for image analysis.

Radioimmunoprecipitation: BHK-21 cells, grown to 75% confluency, were infected

for 1 h at 37oC with EAV at a MOI of 1. Cells were washed once with PBS and incubated

at 37o

C in fresh DMEM. At 48 h post-infection, the culture medium was removed and

cells were starved for 1 h in DMEM without methionine. Then, cells were labeled for 5 h

with 50 μCi of EasyTag EXPRESS protein-labeling mix ([35S]methionine and

[35S]cysteine; specific activity, 407 MBq/ml; Perkin-Elmer). After 5 h, cells were

harvested, washed twice with cold PBS, and lysed in RIPA buffer (1% Triton X-100, 1%

sodium deoxycholate, 150 mM NaCl, 50 mM Tris-HCl [pH 7.4], 10 mM EDTA, 0.1%

sodium dodecyl sulfate [SDS]) containing 1 mM phenylmethylsulfonyl fluoride (PMSF).

Cell lysates were incubated on ice for 20 min, centrifuged at 14,000 RPM for 30 min in a

microcentrifuge and the supernatants were put aside for immunoprecipitation (IP). For IP,

cell lysates were adjusted with RIPA buffer to a final volume of 100 l, and incubated

with 1 l of 3E2 MAb against EAV N for 2 h at room temperature. The immune

complexes were incubated with 7 mg of protein A-Sepharose CL-4B beads (Amersham

Biosciences) for 16 h at 4°C with constant agitation. In the following day, the complexes

were centrifuged at 6,000 RPM for 2 min, and beads were washed with RIPA buffer two

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times and resuspended in 30 l of SDS-polyacrylamide gel electrophoresis (PAGE)

sample buffer (10 mM Tris- HCl [pH 6.8], 25% glycerol, 10% SDS, 0.12% [wt/vol]

bromophenol blue) containing 10% -mercaptoethanol. The mixtures were boiled for 5

min and run on 12% SDS–PAGE gel. The gels were dried and analyzed by a

Phosphorimager (model SI; Molecular Dynamics).

RESULTS

Subcellular localization of EAV N. To study the nucleolar localization of EAV N,

BHK-21 cells were infected with EAV and treated with and without LMB. Cells were

fixed with methanol and stained with a monoclonal antibody against EAV N as well as

DAPI at different time points post LMB treatment. The results of time course study of

EAV infection in BHK-21 cells are shown in Fig. 3.1. As demonstrated in Fig. 3.1, the

EAV N protein localizes in the nucleolus as early as 8 h post-infection (p.i). In the

absence of LMB, many cells showed the nucleolar localization until 12 h p.i. However,

during later stages of infection, the nucleolar localization of EAV N was limited to only a

few cells. At 24 h p.i., the EAV N localization was observed mostly in the cytoplasm and

at 36 h p.i., most of the infected cells were positive for cytoplasmic staining of EAV N

and no nucleolar or nuclear localization was observed (Fig. 3.1, panel A). In the presence

of LMB, EAV N was localized into nucleolus at early and late phases of infection (Fig.

3.1, panel B).

The EAV nucleocapsid protein was expressed as an EGFP-N fusion protein in

HeLa cells and examined for subcellular distribution. Cells were counterstained with the

nuclear stain, DAPI (Fig 3.2). Localization of the PRRSV N protein in infected MARC-

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Fig 3.1. Localization of EAV N in the nucleus and nucleolus of BHK-21 cells at different

time points. Cells were fixed at the indicated times and stained with EAV N specific

MAb followed by staining with goat anti-mouse Ab conjugated with AlexaFluor 488.

Cells were then visualized by fluorescence microscopy. Panels demonstrate EAV-

infected BHK in the absence (A) or presence (B) of leptomycin B (LMB).

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Fig 3.2. Subcellular localization of EAV N protein. HeLa cells grown on the coverslips

were transfected with EGFP-EAV N fusion protein genes using Lipofectin. At 24 hrs

post-transfection, cells were fixed with methanol and stained with DAPI for nuclear

staining. PRRSV-infected MARC-145 cells were stained with MAb SDOW-17, followed

by staining with goat anti-mouse Ab conjugated with AlexaFluor 488 and DAPI for

nuclear staining. Cells were visualized by fluorescence microscopy. Panels A, HeLa

without transfection; B, PRRSV-infected MARC-145 cells stained with the N-specific

MAb SDOW-17; C and D, EAV-infected BHK-21 cells stained with EAV N protein

specific MAb in the absence (C) or presence (D) of leptomycin B respectively; E, HeLa

cells with EAV N-EGFP; F, HeLa cells with EGFP-EAVN. Arrows show the nucleolar

staining of the N protein.

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145 cells, is shown for the purpose of comparison (Fig. 3.2B). Staining of PRRSV N

showed the accumulation of N in the nucleus. Similarly, both EAV N-EGFP and EGFP-

EAV N fusion proteins showed the accumulation of fluorescence in the nucleus with a

distribution pattern similar to PRRS N (Fig. 3.2E and F, respectively). These results

indicate that EAV N was localized both in the cytoplasm and the nucleolus.

Identification and mapping of NLS in the EAV N. Sequence analysis of the EAV N

protein identified the presence of a basic region, which resembles a ‘bipartite’ type NLS

in the EAV N protein predicted by the PSORT II program (Nakai and Kanehisa, 1992).

This motif was located at positions 4-20 as 4-RRSRPQAASFRNGRRRQ-20 (Fig. 3.3A).

To determine the functional significance of this NLS-like sequence, a series of deletion

mutants were made by progressively deleting amino acids from the C- and N-termini of

EAV N (Fig. 3.3B). The mutant constructs were then transfected into HeLa cells and

their fluorescence was examined. EAVN C40-EGFP, EAVN C70-EGFP, and EAVN

C87-EGFP showed fluorescence in the nucleus and nucleolus (Fig. 3.3C, panels B, C and

D). In contrast, EAVN N20-EGFP in which the putative motif was deleted showed the

absence of nuclear fluorescence but the presence of cytoplasmic fluorescence (Fig. 3.3C,

panel D). These results indicate that the predicted ‘bipartite’ sequence may be a

functional NLS that translocates EAV N to the nucleus and nucleolus in transfected cells.

To confirm the function of the predicted NLS and to identify core amino acids essential

for this function, individual arginine (R) residues of 4-RRSRPQAASFRNGRRRQ-20

were substituted with glycine (G). Double, triple, quintuple, sextuple, and septuple G

mutants of EAV N were constructed (Fig 3.4). Individual mutants were expressed in

HeLa cells and visualized by fluorescence microscopy (Fig. 3.4). All of mutants with R

84

85

Fig 3.3. Identification of NLS in EAV N protein and its functional mapping. (A)

identification of a putative ‘bipartite’ type NLS in the EAV N protein predicted by the

PSORT II program (Nakai and Kanehisa, 1992). The PRRSV N protein sequence is

aligned for comparisons. Dashes indicate amino acid deletions. Boxes indicate ‘bipartite’

(for EAV), ‘pat-4’ and ‘pat-7’ (for PRRSV) type NLS sequences. (B). Schematic

presentation of EAVN deletion mutants fused with EGFP. The darkened areas represent

predicted NLS at amino acid positions 4-20. (C). Subcellular localization of EAV N

mutants. Panels A, wild type EAVN-EGFP; B-E, deletion construct: EAVN-EGFP-C40

(B); EAVN-EGFP-C70 (C); EAVN-EGFP-C87 (D); EAVN-EGFP-N20 (E).

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Fig 3.4. Subcellular localization of EAV N NLS mutants. Names of mutants and their

amino acid sequences are indicated. Panels show immunofluorescence of each mutant .

Overlay, superimposition with DAPI staining.

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Fig 3.5. Interaction of EAV-N with importin proteins by GST-pull down assay. Panels A

represents GST pull-down assay using EAV-infected BHK-21 cell lysates and panel B

represents the assay using in vitro translated product. Lane 1 is the immunoprecipitation

(IP) of EAV-infected cells with EAVN MAb. Lanes: 2 and 8, GST; 3 and 9, GST-Jenv as

a negative control; 4 and 10, GST-Imp-α; 5 and 11, GST-Imp-β; 6 and 12, GST-EAVN;

7, uninfected BHK-21 cell lysates; 13 in vitro translated EAV N.

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residues substituted with G showed nuclear localization (Fig. 3.4). This suggests that

these residues may not be the key amino acids to the nuclear localization of EAV N.

Molecular basis for EAV N nuclear localization. To further determine the basis for

EAV N nuclear localization, the interactions between importin proteins and EAV N were

examined. Importin-α and importin-β were expressed in E. coli as a GST-fusion protein

and coupled to glutathione-sepharose beads. The EAV N protein was synthesized either

in vitro by transcription and translation or prepared from EAV infected BHK-21 cells in

the presence of [35

S]methionine. Importin-coupled GST beads were then incubated with

the radiolabelled EAV N protein, and the bound proteins were resolved by SDS-PAGE

and visualized by autoradiography. Despite some degree of non-specific binding of EAV

N to GST, it seems that EAV N interacts with both GST-Imp-α, GST-Imp-β in both in

vitro and cells by pull down assay (Fig 3.5 A and B).

Subcellular localization of SHFV N. HeLa and Cos-1 cells were grown on coverslips

and transfected with genes encoding EGFP-SHFV N and SHFV N-EGFP fusion proteins

using Lipofectin. At 24 hrs post-transfection, cells were fixed with methanol and stained

with DAPI for nuclear staining followed by fluorescence microscopy. The results indicate

that in both cell types, SHFV N-EGFP and SHFV N-EGFP were localized in the nucleus

and nucleolus of transfected cells (Fig. 3.6).

Identification and mapping of NLS in the SHFV N protein. To identify an NLS in

SHFV N protein the PSORT II program (Nakai and Kanehisa, 1992) was used and a ‘pat-

7’ type NLS was predicted at amino acid positions 22-28 of the N protein (Fig. 3.7A). In

order to examine the functional significance of the ‘pat7’ NLS, a series of deletion

mutants of SHFV N were constructed as EGFP fusion proteins (Fig. 3.7B). HeLa cells

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Fig. 3.6. Subcellular localization of SHFV-N protein in HeLa and Cos-1 cells. Cells

grown on the coverslips were transfected with EGFP-SHFV N fusion protein genes using

Lipofectin. At 24 hrs post-transfection, cells were fixed with methanol and stained with

DAPI for nuclear staining. Cells were then visualized by fluorescence microscopy (model

AX70, Olympus). Panels A, Cos-1 cells with SHFV N-EGFP; B, HeLa cells with SHFV

N-EGFP; C, Cos-1 cells with EGFP-SHFV N; D, HeLa with EGFP-SHFV N.

90

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Fig 3.7. Identification of NLS motif in SHFV N protein. (A) a ‘pat-7’ type NLS in the

SHFV N protein predicted by the PSORT II program (Nakai and Kanehisa, 1992).

PRRSV N protein sequence is aligned for comparison. Dashes indicate amino acid

deletions. Boxes indicate ‘pat-7’ (for SHFV) and ‘pat-7’ (for PRRSV) type NLS

sequences. (B) Schematic presentation of SHFVN deletion mutants fused with EGFP.

Darkened areas represent predicted NLS at amino acid positions 22-28. (C) Subcellular

localization of SHFV N mutants. Panels A, wild type SHFVN-EGFP; B, Deletion

construct SHFVN-EGFP-C20; C, deletion construct SHFVN-EGFP-C40; D, deletion

construct SHFVN-EGFP-C70; E, deletion construct SHFVN-EGFP-N30.

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were transfected with the mutant constructs and subcellular localization of SHFV N

mutant proteins was determined (Fig. 3.7C). The SHFV N constructs containing the

putative ‘pat7’ type of NLS localized in the both cytoplasm and nucleolus of gene-

transfected HeLa cells (Fig. 3.7C, panels B, C, and D), whereas the mutants that did not

contain the predicted NLS showed only cytoplasmic staining (Fig. 3.7C, panel E). These

observations indicate that the putative NLS identified at positions 22-28 was a functional

motif. To determine core amino acids essential for the function of ‘pat7’ type NLS of

SHFV N, the R residues in NLS (22-PQRPRRS) of SHFV N were individually mutated

to G (Table 3.3). The NLS-null SHFV N gene was transfected to HeLa cells followed by

staining with DAPI and fluorescence microscopy. Fig. 3.8 demonstrates the subcellular

location of the NLS-null SHFVN-EGFP mutant in the transfected HeLa cells. The NLS-

null mutant protein of SHFV N showed neither nuclear nor nucleolar localization, and

fluorescence was seen only in the cytoplasm of transfected cells. This indicates that all

arginine residues of SHFV N protein are important for its nuclear localization (Fig.

3.8A).

Molecular basis for SHFV N nuclear localization. To determine the molecular basis

for SHFV N nuclear localization, interactions between importin shuttle proteins and the

N protein were examined. The SHFV N protein was synthesized in vitro by transcription

and translation in the presence of [35

S] methionine. The GST fusion proteins of Imp-α

and Imp-β expressed in E. coli and coupled to glutathione-Sepharose beads were then

incubated with the radiolabelled SHFV N protein. After washing, bound proteins were

released by boiling for 5 min in SDS sample buffer and resolved by 12% SDS PAGE for

autoradiography using Phosphorimager. The result shown in Fig. 3.9 indicates that SHFV

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Table 3.3. Site-directed mutagenesis of the SHFV N NLS motif and subcellular

localization of NLS mutants.

Construct Sequence of NLSa

Localizationb

Cy Nu No

SHFV-N 22-PQRPRRS ++++ ++ ++++

SHFV-N- R24,26,27G 22-PQGPGGS-28

++++ - -

aIndividual Arginines (underlined) in the NLS were substituted with glycines.

bFor scoring of the fluorescence signal for each mutant construct, 100 transfected cells

were counted presented as + or -. Each one ‘+’ represents fluorescence of 25 transfected

cells either in the cytoplasm (Cy), nucleus (Nu), or nucleolus (No).

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Fig. 3.8. Subcellular localization of the NLS-null SHFVN-EGFP mutant. The basic

residues in the NLS motif (22-PQRPRRS) of SHFV N were mutated into 22-

PQGPGGS-28. The NLS-null SHFV N gene was transfected to HeLa cells followed by

staining with DAPI and fluorescent microscopy. A, wild type SHFVN-EGFP; B, NLS-

null SHFVN-EGFP.

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Fig 3.9. Interaction of SHFV N with importin proteins by GST-pull down assay.

Importin-α and importin-β-glutathione-S-transferase (GST) fusion proteins were

expressed in E-coli and coupled to glutathion Sepharose 4B beads (Pharmacia). The

beads-bound proteins were incubated with radiolabelled in vitro-translated SHFV N

protein overnight at 4°C with constant agitation. After washing, bound proteins were

released by boiling for 5 min in SDS sample buffer and resolved by 12% SDS PAGE for

autoradiography using Phosphorimager. Lanes: M, protein marker; 1, in vitro translated

product of SHFV N; 2, GST pull-down assay for SHFV N and GST; 3, SHFV N and

GST-Jenv; 4, SHFV N using importin-α, 5, SHFV N and importin-β.

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N interacts with both importin-α and importin-β, with a stronger affinity for importin-α

(Fig. 3.9, lanes 5 and 6).

DISCUSSION

Nuclear localization of capsid protein for cytoplasmic RNA viruses has been

investigated in this chapter using EAV and SHFV as a model. Nuclear localization of

capsid protein from cytoplasmic RNA viruses has been observed previously, and

examples include capsid proteins of Semliki Forest virus, dengue fever virus, and

hepatitis C virus, all of which are translocated to the nucleus and nucleolus (Favre et al.,

1994; Yasui et al., 1998, Wang et al., 2002). The exclusive replication of nidoviruses in

the cytoplasm of infected cells has been previously reported for MHV, which replicates

in enucleated cells (Brayton et. al, 1981). This indicates that these viruses do not need

nucleus for their replication. But, nuclear and nucleolar localization for nucleocapsid

proteins of PRRSV, EAV, IBV, MHV and TGEV has been observed (Hiscox et al, 2001;

Wurm et al, 2001; Rowland et al., 1999; Tijms et al., 2002). The nuclear localization of

proteins is mainly mediated by the nuclear localization signal (Golrich and Kutay, 1999).

In arteriviruses, PRRSV N contains two putative NLS motifs, ‘pat 7’ and ‘pat 4’

(Rowland et al., 2003). The location of “pat 7”, the primary and active NLS of the

protein, is between positions 41-47 (Rowland et al., 2003). The ‘pat-4’ NLS is also

functional but only in the absence of ‘pat7’ motif and leads PRRSV N to the nucleus

(Rowland et al., 2003). In addition to PRRSV N, we have shown study that the LDV N

protein is also localized in the nucleus and nucleolus of transfected cells (Mohammadi et

al., 2009). Our studies using mutant LDV N constructs showed that the nuclear

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localization of LDV N was dependent on a ‘pat-4’ motif at positions 38-41 (Fig. 2.2B;

Fig. 2.3), indicating that this motif was indeed functional. The nuclear localization of

nucleocapsid proteins of two other members of the family Arteriviridae, EAV N and

SHFV N, had not been investigated in detail prior to the current study, even though one

report showed the nucleolar translocation of EAV N in virus- infected cells (Tijms et al.,

2002). No such information was available for SHFV N. Thus, we conducted detailed

studies and report the nuclear and nucleolar localization of nucleocapsid proteins for both

EAV and SHFV.

EAV N was previously observed to localize in the nucleolus of infected cells

(Tijms et al., 2002). We expanded the study to further examine its nuclear translocation

and identify the presence of an NLS. Based on our data, EAV N is localized in the

nucleolus of infected cells at the early stage of virus infection, as the fluorescence was

detected at 8 hr p.i (Fig. 3.1). EAV N did not translocate to the nucleus in the absence of

the first 20 residues at the N-terminus of the protein. This motif contains a putative

bipartite NLS motif (Fig. 3.3. B and C). Fine mapping studies of the putative NLS

showed that the substitution of basic amino acids in this motif did not prevent the nuclear

localization of EAV N (Fig. 3.4). This suggests that EAV N may require all of the N

terminal 20-amino-acids within the motif to be functional for nuclear translocation. A

bipartite NLS has been identified in many eukaryotic proteins such as nucleoplasmin,

human pRB, human p53, and human interleukin 5 (IL5) (Munoz-Fontedla et al, 2003).

Functional bipartite NLSs have also been identified in viral proteins. For example,

nucleoproteins of Thogoto virus and influenza A virus, phosphoprotein of Borna disease

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virus, and the core protein of hepatitis C virus all contain functional bipartite NLSs

(Weber et. al, 1998; Schwemmle et. al, 1999; Suzuki et al., 2005).

We also investigated the nuclear localization of SHFV N. For this protein, a ‘pat-

7’ type NLS was identified at amino acid positions 22-28 (Fig. 3.7). Studies using a series

of deletion mutants of SHFV N showed that the putative ‘pat7’ type of NLS was

necessary to lead SHFV N to the nucleus in the N gene-transfected cells (Fig. 3.7, B and

C). The key amino acids for the NLS of SHFV N were further identified by substituting

arginines in the putative NLS with glycines (Table 3.2). The results showed that all of the

arginines within the motif were essential for nuclear localization of SHFV N (Fig. 3.8A).

The cellular mechanism for nuclear localization of a protein starts by recruiting a family

of cellular proteins, and among them importins are the most common proteins. The

process begins by binding of importin-α to NLS on the cargo protein followed by binding

of importin-β to the importin-α/cargo protein complex, which finally transports the entire

complex to nuclear pore complex (NPC) on the nuclear membrane in the presence of

Ran-GDP. We showed here that EAV N and SHFV N both could bind to importin-α and

importin β (Fig. 3.5 and Fig. 3.9). EAV N appeared to bind to GST as well, but perhaps

this interaction would be non-specific. PRRSV N and LDV N bind to importin-α and

importin-β (Rowland et al., 2003; Mohammadi et al., 2009). Several viral proteins

have been reported to interact with both importin-α and -β. For instance, the

polyomavirus capsid protein requires the interaction with both importin-α and -β through

its NLS to enter the nucleus (Qu et al., 2004), and SV40 VP3 also bins to importinα-α/β

complex to enter the nucleus (Nakanashi et al, 2002). The HIV Rev protein directly binds

to both importin-α and -β, but its binding to importin-α is NLS-independent while

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binding to importin-β is NLS-dependent via its arginine-rich NLS (Henderson et al.,

1997). It is possible that these proteins use another cellular protein which mediates the

transport to the nucleus. Binding of EAV N to GST in addition to GST-importin fusion

proteins could be a source of non-specific binding of the N protein to these proteins.

Since N protein of all arteriviruses are localized to the nucleus and nucleolus, the

nuclear role of N during infection is of major interest. The core protein of hepatitis C

virus is believed to modify cellular functions by preventing translocation of host cell

proteins to the nucleus (Isoyama et al., 2002). The N protein of TGEV has been

suggested to play a role in the disruption of cell division (Wurm et al., 2001). For PRRSV

N, disruption of NLS does not affect virus replication in cell culture, but in pigs, NLS-

null viruses were attenuated (Lee et al., 2006a, 2006b; Pei et al., 2008). A recent study

showed that the nuclear import of PRRSV N from the cytoplasm was faster than the

export from the nucleus and the N protein was not kept in the nucleolus (You et al.,

2008). We also noticed that EAV N protein localizes in the nucleus at the early stage of

EAV infection. Our observation along with another report on PRRSV N (You et al.,

2008) suggest an important role for N during the early stage of PRRSV and EAV

replication in which N is localized predominantly in the nucleolus. N protein of several

member viruses in the order Nidovirales have been shown to co-localize and interact with

fibrillarin and nucleolin in the nucleolus, supporting the hypothesis that N plays a role

during host cell ribosomal biogenesis and cell cycle modulation (Yoo et al., 2003; Chen

et al., 2002).

In summary, we show that EAV N and SHFV N are both localized in the nucleus

and nucleolus of N gene transfected cells. EAV N localization is dependent on a part of

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the protein which contains a bipartite NLS. Also SHFV N has a functional ‘pat7’ type

NLS at positions 22-28. The capsid protein nuclear localization is a common feature in

the Arteriviridae family, and this protein may play a role in replication and pathogenesis

of this group of viruses.

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CHAPTER 4

The Role of Nucleocapsid Protein of Arteriviruses in Cell Cycle Modulation

102

ABSTRACT

Replication of porcine reproductive and respiratory syndrome virus (PRRSV) and

equine arteritis virus (EAV) in the dividing MARC-145 and BHK-21 cells, respectively,

led to accumulation of cells at G2/M phase of the cell cycle. The current study was

conducted to identify the viral component involved in this function. The N protein was

the most probable viral protein for this activity since it localizes to the nucleolus of virus-

infected and N gene transfected cells and interacts with nucleolar proteins including

fibrillarin which participates in the ribosome biogenesis of cells. In BHK-21 cells

expressing GFP-N of arteriviruses (GFP-PRRSV N, GFP-EAV N, GFP-LDV N, and

GFP-SHFV N), the number of cells in G2/M phase was higher compared to untransfected

cell controls. This suggests that the nucleocapsid protein of arteriviruses plays a

modulatory role during cell cycle. The effect of N protein was similar for the all four N

proteins but some difference were observed. PRRSV N and EAV N had more cells at

G2/M stage compared to LDV N and SHFV N. This study for the first time suggests that

N protein of arteriviruses affects cell cycle process in the N-expressing cells.

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INTRODUCTION

The family Arteriviridae is a group of RNA viruses consisting of four different

members: porcine reproductive and respiratory syndrome virus (PRRSV), equine arteritis

virus (EAV), lactate dehydrogenase-elevating virus (LDV), and simian hemorrhagic

fever virus (SHFV) (Snijder and Meulenberg 1998). The Arteriviridae family, along with

the Coronaviridae family, is grouped in the Nidovirales order of viruses (Cavanagh et al,

1997). Despite their replication in the cytoplasm of infected cells, the nucleocapsid (N)

protein of PRRSV and EAV have been shown to localize in the nucleus and nucleolus of

virus-infected and gene-transfected cells (Rowland et al, 1999; Tijms et al, 2002). The N

protein of these viruses is encoded by the 3’ ultimate open reading frame (ORF) of viral

genome and varies in size between 14-15 kDa (Snijder et al, 1998). The N protein of

other member viruses localizes in the nucleus and nucleolus of infected and N-gene

transfected cells (Mohammadi et al., 2009; Chapter 2). Studies have also shown the

colocalization and interaction of PRRSV N with nucleolar proteins fibrillarin and

nucleolin. These proteins are known to be involved in a variety of cellular processes,

including cell cycle progression (Yoo et al., 2003; Chen et al., 2002; Wurm et al., 2001).

A recent study showed that PRRSV N interacts and co-localizes with human I-mfa

domain-containing protein (HIC) which is a cellular transcription factor (Song et al,

2009). HIC itself interacts with cyclin T1 which is part of a protein complex controlling

the progression of cell cycle (Young et al, 2003). This suggests that PRRSV N protein

may play a role in cell cycle modulation of the virus infected cells. A role for nuclear

localization of the N protein and the consequence of its interaction with cellular proteins

has not been investigated previously.

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The cell cycle is a complex process controlled by many regulatory proteins which

leads to mitosis and generation of daughter cells. This process is regulated by specific

cellular proteins, namely cyclins and cyclin-dependent kinases (Cdks). The cell cycle is

divided into four phases: G0/G1 (quiescent phase/ Gap1), S (Synthesis phase), G2/M

(Gap2/ Mitosis), and M. Each stage of cell cycle is controlled by specific cyclins and

Cdks. Alteration in any of these regulatory factors can influence or disturb the normal

cell cycle (Schafer, 1998). Several studies have demonstrated the effect of viruses and

viral proteins on cell proliferation and cycle at different stages, mostly G2/M in the case

of RNA viruses. Examples of viruses arresting the cell cycle at G2/M include human

immunodeficiency virus type I (HIV-1), reoviruses, and coronaviruses (Emmet et al,

2005; Zhao et al, 2005 and Davy and Doorbar et al, 2007). Human Parvovirus B19 stops

the cell cycle at the G2 stage and increases the cellular regulatory factors of mitosis step

(Morita et al, 2001). For Nidovirales, several member viruses of the family

Coronaviridae, such as infectious bronchitis virus (IBV), murine hepatitis virus (MHV),

and severe acute respiratory syndrome (SARS) virus, affect cell cycle at different stages.

IBV induces cell cycle arrest at S and G2/M stage in infected cells (Dove et al, 2006b; Li

et al, 2007). However, investigations show that MHV and SARS virus arrest cell cycle at

the stage of G0/G1 (Chen and Makino 2004; Yuan et al, 2006). There is no report on

modulation of cell cycle by Arteriviridae and their viral proteins. Therefore, the

objectives of this study were firstly to investigate the effects of arteriviruses on cell cycle

and secondly to examine arterivirus N protein for possible involvement in this process.

Here we show that PRRSV and EAV also regulate cell cycle in MARC-145 and BHK-21

105

cells, respectively. Further, we demonstrate that N protein is the viral component

associated with this modulation in BHK-21 cells.

MATERIALS AND METHODS

Cells and viruses. Baby hamster kidney cells (BHK-21) and MARC-145 (Kim et al,

1993) which is a derivative line of MA104 cells were used in this study. Cells were

maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10%

heat inactivated fetal bovine serum (FBS, Gibco BRL) in a humidified incubator with 5%

CO2 at 37ºC. EAV strain Bucyrus (Doll et al., 1957) and PRRSV strain PA8 (Wootton et

al., 2000) were grown in MARC-145 cells and BHK-21 cells, respectively.

Plasmid constructs. The pEGFP-N1 (Clontech Inc.) was used in this study. EAV N,

LDV N, and SHFV N coding sequences were sub-cloned into Bam HI site of the

pEGFP-N1 upstream of the enhanced green fluorescence protein (EGFP) in-frame from

PCR products using specific primers (Mohammadi et al., 2009; Chapter 3 Table 3.1 and

3.3) and the fusion genes EAV N-GFP, LDV N-GFP, and SHFV N-GFP were

constructed. To amplify the EAV N gene, pEAV030 containing the full genome sequence

of EAV (van Dinten et al., 1997) was used as the template for PCR reaction. SHFV N

gene was cloned in pGEM-T using a cDNA library of SHFV genome. It was then

amplified by PCR from the plasmid using specific primers and sub-cloned upstream of

EGFP in the pEGFP-N1 and in frame. The LDV-N gene was amplified by PCR from a

plasmid containing the LDV genomic fragment, and using the Bam HI sequence at both

ends of the primers, cloned into pEGFP-N1 upstream of the EGFP gene, respectively.

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The PRRSV N-EGFP fusion protein has been described previously (Rowland et al., 1999;

Rowland et al., 2003).

Co-staining of viral N and fibrillarin. For immunofluorescence purpose, HeLa cells

were used to express LDV N-EGFP fusion protein and BHK-21 cells were used for EAV

infection. Cells were seeded on coverslips in 35 mm diameter dishes. At 75% confluency,

cells were either transfected with 2 μg of a plasmid containing LDV N-EGFP fusion

construct and Lipofectin (Invitrogen) according to the manufacturer’s instruction or

infected with EAV at a multiplicity of infection (MOI) of 1. After 24 hours, cells were

fixed by methanol, and stained with a specific antibody for fibrillarin followed by

staining with goat anti-rabbit IgG conjugated with Texas red dye (Molecular Probes,

Eugene, Oreg.,) or Alexa fluor 488, and finally stained with DAPI (4',6-Diamidino-2-

phenylindole dihydrochloride). The EAV infected cells were co-stained with a mAb

against EAV N (MAb 3E2, IgG isotype) which was a generous gift from Dr. Udeni B.R.

Balasuriya (Department of Veterinary Science, Gluck Equine Research Center,

University of Kentucky, Lexington, USA) and fibrillarin followed by incubation with

goat anti-mouse immunoglobulin G conjugated with AlexaFluor 488 (Molecular Probes)

and goat anti-rabbit immunoglobulin G conjugated with Texas red dye (Molecular

Probes, Eugene, Oreg.). After washing, cells were finally stained with DAPI. After last

staining, cells were washed and mounted on a slide using 40 μl of the mounting buffer

(60% glycerol, 0.1% sodium azide in PBS), and visualized for fluorescence.

Flow cytometry of virus infected cells. Fig. 4.1 shows the experimental design used for

analysis of arteriviruses and their nucleocapsid protein on cell cycle. 06 cells of MARC-

145 or BHK-21 were seeded onto every 100 mm diameter cell culture dishes. The

107

Fig 4.1. Experimental design for analysis of arteriviruses and their nucleocapsid protein

on cell cycle.

108

following day, cells were either mock or virus-infected at a multiplicity of infection of 5,

with PRRSV or EAV for MARC-145 and BHK-21 cells, respectively. At different time

points post-infection (12, 24, and 48 hrs), cell monolayers were washed, trypsinized, and

fixed in 4% paraformaldehyde (PFA) for 10 min at room temperature. Then, cells were

washed twice with cold PBS and stained with propidium iodide (PI) (50g/ml propidium

iodide, 0.1% NP-40, 0.1% Sodium citrate, and 20 g/ml RNase A in PBS). After 30 min

incubation at room temperature, the DNA content of 30,000 cells was assessed using

FACScan flow cytometry instrument (Becton Dickinson). Then DNA content of the

acquired cells was analyzed using the ModFit LT 3.0 DNA analysis software. Each

experiment was performed in triplicate and repeated three times.

Flow cytometry of N gene transfected cells. Approximately 3x10

6 BHK-21 cells per

dish were seeded onto 100 mm diameter dishes. After 24 h, cells were transfected with

GFP-N gene using 60 l of 1 mg/ml of stock solution of PEI (Linear polyethylenimines;

Polysciences Inc.) and 20 ug of plasmid DNA in serum-free DMEM. At 16 hrs post-

transfection, the medium was replaced with DMEM containing 10% fetal bovine serum.

At 60 h post-transfection, cells were harvested, fixed in 4% paraformaldehyde and

stained with PI for 30 minutes at room temperature. Then the DNA content of 10,000

transfected cells was analyzed by FACScan.

RESULTS

Co-localization of LDV N and EAV N with fibrillarin. Previously, co-localization of

PRRSV N and the nucleolar protein fibrillarin was reported (Yoo et al., 2003). In this

study, this feature was examined for two other member viruses of the Arteriviridae

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Fig 4.2. Co-localization of LDV N and EAV N with fibrillarin. A) HeLa cells were

transfected with LDV N-EGF using Lipofectin. After 24 hours, cells were fixed by

methanol and stained with an antibody against fibrillarin followed by staining with goat

anti-rabbit IgG conjugated with Texas red dye and finally stained with DAPI and

visualized by immunofluorescence microscopy. B) EAV infected BHK-21 cells. BHK-21

cells were seeded on coverslips in 35 mm diameter dishes. At confluency of 70% cells

were infected with EAV at a MOI of 1. The EAV infected cells were co-stained with an

antibody against EAV N (MAb 3E2) and fibrillarin followed by incubation with goat

anti-mouse immunoglobulin G conjugated with AlexaFluor 488 (Molecular Probes) and

goat anti-rabbit immunoglobulin G conjugated with Texas red dye. After washing cells

were finally stained with DAPI and mounted on a slide, and visualized for fluorescence.

Arrows indicate the localization of the N protein and fibrillarin. Cy-N and No-N show the

cytoplasmic and nucleolar localized nucleocapsid proteins with the green fluorescence.

Fib is fibrillarin with the red fluorescence. Fib+N shows the co-localization of N protein

with fibrillarin.

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family to determine if it was a common feature of N protein in this group of viruses. LDV

N- EGFP fusion protein was expressed in HeLa cells and stained with a rabbit polyclonal

antibody against fibrillarin (Abcam, ab5821) as well as with DAPI. The result in Fig. 4.2

A shows that LDV N co-localizes with fibrillarin in the nucleolus of the N-gene

transfected cells. For EAV, BHK-21 cells were infected and the N protein was examined

by staining. It was apparent that EAV N also co-localized with fibrillarin in virus-infected

BHK-21 cells (Fig. 4.2B).

PRRSV infection of asynchronized MARC-145 cells increases the number of cells in

the G2/M phase of cell cycle. Co-localization of N protein with nucleolar proteins

suggests that N protein may modulate the normal cellular activities such as cell cycle

progression. This possibility was examined first using virus-infected MARC-145 cells

infected with PRRSV at an MOI of 5 and were harvested at 12, 24, and 48 hr p.i. Cells

were then stained with PI and DNA contents of cells were analyzed by flow cytometry.

To monitor the infection, cytopathic effects (CPEs) were also determined using

microscopy. CPE was increased in virus-infected cells and the increase was time-

dependent (Fig. 4.3). Fig. 4.4 shows the cell cycle profile of PRRSV-infected MARC-145

cells. The number of virus-infected cells was significantly (P 0.05) higher compared to

mock-infected (uninfected) cells in the G2/M phase at 12 and 24 h p.i. At 48 hrs, the

number of infected cells was higher in the S phase compared to mock-infected cells.

Accumulation of EAV infected BHK-21 cells in the G2/M phase. To examine if the

cell cycle modification was common for arteriviruses, EAV was chosen. BHK-21 cells

were either mock-infected or virus-infected with EAV (strain Bucyrus) at a MOI of 5. At

24 h p.i, cells were harvested and stained with PI as described for DNA quantification.

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Fig 4.3. CPE of PRRSV-infected MARC-145 cell at different time points. Panels A, B,

and C are mock-infected cells at 12h, 24h, and 48h post infection respectively and panels

D, E, and F demonstrate the PRRSV infected MARC-145 cells (with an MOI of 5) at

12h, 24h, and 48h post-infection with PRRSV. Arrows show the CPE.

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Fig. 4.4. Cell cycle profile of PRRSV-infected MARC-145 cells. Averaged values from

three independent experiments are presented. MARC-145 cells were infected with

PRRSV strain PA8 at an MOI of 5. At 12hr, 24hr, and 48hr post-infection, cells were

trypsinized, washed, fixed with 4% PFA, and stained with PI. After 30 min, the amount

of DNA of cells were analyzed using flow cytometry. The upper graph demonstrates the

comparison of mock- and PRRSV-infected cells at different stages of cell cycle. The

asterisks indicate significant difference between two groups (P-value less than 0.05) and

the error bars show the standard error. The lower graph shows the trend of cell cycle

analysis at different time points in mock- and PRRSV- infected cells separately.

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Fig. 4.5. Cell cycle profile of EAV-infected BHK-21 cells. BHK-21 cells were infected

with EAV strain Bucyrus at an MOI of 5. At 24 hrs post-infection cells were trypsinized,

washed, fixed with 4% PFA, and stained with PI. After 30 min, the amount of DNA of

cells was analyzed using flow cytometry . Each experiment is an average of three

independent experiments. The upper graph demonstrates the comparison of mock- and

EAV-infected cells at different stage of cell cycle. Asterisks demonstrate a significant

difference between groups (the P-value less than 0.05) and the error bars are

representative of standard errors. The lower graph shows the trend of cell cycle analysis

at different time points in mock- and EAV-infected cells.

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Fig. 4.5 demonstrates the cell cycle profile of from this experiment. The number of cells

was significantly (P 0.05) higher at the S and G2/M phases of cell cycle in EAV-

infected cells compared to mock-infected cells.

Nucleocapsid protein of arteriviruses arrests cell cycle in G2/M stage. Since PRRSV

and EAV appear to modulate the cell cycle during infection, it was of interest to

determine the viral component responsible for this function. Since the nucleocapsid

protein of these viruses co-localizes with nucleolar proteins, the N protein was examined

for this function. BHK-21 cells were either untransfected or transfected with GFP, GFP-

PRRSV N, GFP-EAV N, GFP-LDV N, GFP-SHFV N, and the patterns of cell cycle were

determined at 60 h post-transfection (Fig. 4.6). The cell cycle profile was similar in

untransfected and GFP-transfected cells. The number of cells in the G2/M phase was

higher in N-gene transfected cells for all members of the Arteriviridae family compared

to the negative controls (untransfected and GFP expressing BHK-21 cells). However, for

PRRSV N and EAV N gene-transfected cells the number of cells in the G2/M phase were

more than those for LDV N and SHFN transfected cells.

DISCUSSION

In this study we have demonstrated that viruses in the Arteriviridae family can

cause infected cells to accumulate in the G2/M stage of the cell cycle. We further show

that the nucleocapsid protein of arteriviruses is responsible for this G2/M accumulation.

PRRSV-infected MARC-145 cells were analyzed for DNA content at different

time points post-infection (p.i.). It was noticed that the number of cells at G2/M was

significantly higher in PRRSV-infected cells compared to uninfected cells at 12 and 24 h

115

Fig. 4.6. Cell cycle profiles in N-gene transfected BHK-21cells. BHK-21 cells were

transfected with GFP, GFP-PRRSV N, GFP-EAV N, GFP-LDV N, and GFP-SHFV N.

At 60 hrs post-transfection cells were trypsinized, washed, fixed with 4% PFA, and

stained with PI. After 30 min, the amount of DNA was analyzed using FACScan.

116

p.i. (Fig. 4.4). However at 48 h p.i at which the maximum CPE was observed (Fig 4.3F),

the number of cells at G2/M in the infected group was lower than that of uninfected cells.

At 48 h p.i. infected cells were significantly lower and higher at the G0/G1 phase and the

S phase of cell cycle respectively. It seems that infected cells move faster to G2/M at 24

hours and then to S phase at 48 h p.i. in comparison with uninfected cells. This indicates

that virus infected cells appear to cycle faster than uninfected ones, which can be

considered as favored for virus replication. EAV infection of BHK-21 cells led to a

significant increase in the number of cells at the S and G2/M phases and a significant

decrease in the number of cells at the G0/G1 phases of cell cycle at 24 h p.i (Fig. 4.5).

In order to determine which viral components are involved in cell cycle

modulation by arteriviruses, the most likely candidate was the N protein as it interacts

with nucleolar proteins involved in the cell cycle (Fig 4.2). As it is shown in Fig 4.2, both

LDV N and EAV N proteins colocalize with fibrillarin, a host cell nucleolar protein.

These data along with the previous findings of PRRSV N co-localization with fibrillarin

indicate that co-localization of N with fibrillarin is a common feature in arteriviruses,

suggesting that N protein may be involved in modulation of the cell cycle. We

determined if N had any impact on cell cycle modification. N proteins of arteriviruses

were transfected in to BHK-21 cells as GFP fusion proteins and the DNA content of the

transfected cells was analyzed. Our results show that the number of cells at the G2/M

stage was higher at 60 h post-transfection in the cells expressing N proteins of the

Arteriviridae family compared to untransfected and GFP transfected negative controls

(Fig 4.6). However, the PRRSV N and EAV N had stronger activity on the number of

cells at G2/M compared to LDV N and SHFV N at 60 h post transfection (Fig. 4.6). Our

117

results indicate that the N proteins of arteriviruses modulate the cell cycle in the similar

manner but they may function differently.

Cell cycle modification by RNA viruses has been reported, particularly with

regards to stalling at G2/M phase (Davy and Doorbar, 2007; Zhao and Elder 2005). The

stage of cell cycle might be a determinant of permissiveness and level of virus production

for some viruses. For instance, Dengue virus type 2 infects different types of permissive

cells based on the cell cycle stage. HepG2 cells are more permissive to Dengue virus

infection as well as virus production at the G2/M step of cell cycle (Phoolcharoen and

Smith. 2004).

The involvement of viral proteins in modulation of the cell cycle has been studied

for several viruses. In this regard, mostly non-structural (NS) viral proteins are involved

in cell cycle modulation. The Vpr protein of HIV-1, which is a nucleo-cytoplasmic

protein, has been found to arrest cells in the G2/M phase of the cell cycle (Mahalingam et

al, 1997; Matsuda et al, 2003). Furthermore, the σ1S protein of alpha reovirus also stops

cell cycle at the G2/M phase (Poggioli et al, 2000). The paramyxovirus simian virus 5

(SV5) V protein slows down the cell cycle process (Lin and Lamb 2000). Hepatitis C

virus non-structural proteins 5A and 2 have been shown to regulate cell cycle phases

(Ghosh et al, 1999; Yang et al, 2006). The NS protein p28 of MHV and NS protein 7a of

SARS coronavirus are shown to block the G0/G1 progress (Chen et al, 2004, Yuan et al,

2006). Structural proteins of RNA viruses can also play a role in cell cycle modulation.

Transmissible gastroenteritis virus nucleocapsid protein (TGEV N) has been shown to

modify the cell cycle. When cells were transfected with TGEV N, only 30% of

transfected cells were dividing, which suggested cell cycle arrest at the G2 phase (Wurm

118

et al, 2001). The nuclear localization of IBV N protein is dependant on stage of the cell

cycle. In G2/M synchronized Vero cells expressing IBV N, the nuclear localization of N

was monitored by live microscopy and the percentage of nuclear localization was found

to be higher. This indicates that at the G2/M stage of cell cycle is more suitable for

nuclear localization of IBV N (Cawood et al, 2007).

Here we showed that the N protein affects cell cycle by accumulation of cells at

G2/M stage. These findings suggest that the N protein plays a significant role in

pathogenesis of these viruses, and further studies are needed to scrutinize its mechanism

of function in host cells and pathogenesis of arteriviruses.

119

CHAPTER 5

General Discussion

Nuclear localization of the most abundant structural protein of arteriviruses,

which is N protein, in the virus infected cells has been of great interest to investigators

working on these viruses. Moreover, the exclusive cytoplasmic replication of

arteriviruses with no need for cell nucleus makes this phenomenon more interesting. The

primary objective of this research was to investigate the nuclear localization property of

the nucleocapsid (N) protein of the viruses in the family Arteriviridae. We also

investigated the possible role of N protein in modulation of the host cell function in virus-

infected cells and N gene-transfected cells.

The nucleolar localization for N protein in the family of Arteriviridae was first

reported for PRRSV N. The presence of a functional NLS in PRRSV N as well as the

mechanism for its nuclear translocation, have been investigated thoroughly. However,

there was little information available for N proteins of other members of the family

including LDV, SHFV, and EAV. Since LDV exhibits limited cell tropism and grows

only in a limited number of mouse primary macrophages, its subcellular localization was

investigated using the cloned LDV N gene. Subcellular localization of LDV-N was

determined by tagging N protein with enhanced green fluorescence protein (EGFP) on

the N- or C-terminus. The fusion proteins localized to the nucleus and nucleolus of gene-

transfected cells. Further studies using a series of mutant constructs of LDV N indicated

that a functional ‘pat4’ type NLS consisting of KKKK is involved in nuclear localization

of this protein and the key amino acids for this activity were identified. The location of

NLS for LDV N was in the similar region of PRRSV N functional NLS. Furthermore, the

120

LDV N interaction with the importin-α and -β proteins suggests that its nuclear

localization occurs through the importin-mediated nuclear transport pathway. These

findings show that the nuclear localization is the property of LDV N in addition to

PRRSV N.

Despite the report on nucleolar localization of EAV N in infected BHK-21 cells

(Tijms et al., 2001), little was known about the details of its nuclear localization. Thus,

we conducted further studies on EAV N nucleolar localization. EAV N fusion constructs

were made to fuse with EGFP on the N- and C-termini and their cellular distribution was

investigated in the gene-transfected cells. Both N-EGFP and EGFP-N fusion proteins of

EAV N showed accumulation of fluorescence in the nucleus and nucleolus of gene-

transfected cells. The presence of a putative ‘bipartite’ type NLS on EAV N sequence at

amino acids position 4-20 was found using the PSORT II program. Based on the cellular

distribution of a series of deletion mutants of EAV N , it appeared that EAV N needed the

entire motif of 4-20 residues for nuclear localization. Because of not clear interaction of

EAV N with importin-α and -β proteins, it is possible that it may not utilize the importin-

regulated nuclear transport pathway. Its interaction with GST itself and an unrelated

GST-fusion protein might be a nonspecific binding. The other possibility could be that it

uses other cellular protein to go to nucleus.

The N protein of SHFV is the least characterized N protein in the family so far.

The SHFV N gene was cloned as a fusion protein with EGFP on the N- or C-terminus.

Both SHFV N-EGFP and EGFP-SHFV N fusion proteins showed nuclear and nucleolar

localization. The significance of a functional ‘pat-7’ type NLS at amino acid positions

22-28 predicted by PSORT II, was determined by constructing a group of progressively

121

deleted mutants for SHFV N protein. Furthermore, the interaction of SHFV N protein

with importin-α suggests that it recruits this chaperon protein for its nuclear transport.

These findings resemble the data for the N proteins of other members of arteriviruses,

indicating that nucleolar localization of the N protein is a common and conserved feature

for Arteriviridae.

The conserved feature of nucleolar localization of N protein among all members

of the Arteriviridae family suggests that it plays an important accessory role during viral

replication in addition to its participation in virion assembly. The nuclear localization of

nucleocapsid protein of PRRSV was investigated in vitro intensively and only one in vivo

study was conducted using infectious clone harboring the PRRSV N with deleted NLS.

This study showed that the mutant protein had limited replication compared to the wild

type PRRSV which indicates that N protein is important in the pathogenesis of PRRSV

(Lee et al, 2006a). PRRSV N, like N protein of the member viruses of Coronaviridae,

interacts with nucleolar and cellular proteins involved in regulating cell cycle. The co-

localization of LDV N and EAV N proteins with fibrillarin indicates that it is also a

shared property between members of Arteriviridae. Since no data was available for the

role of N in cell cycle modulation by Arteriviridae, the research on this subject would

elucidate a very important aspect of arterivirus-cell correlation. Our results showed that

replication of PRRSV and EAV in growing cells leads to accumulation of cells at the

G2/M phase of the cell cycle. We further examined if N protein was the viral component

involved in this function. The N protein of all 4 member viruses of the Arteriviridae

family was so expressed in BHK-21 cells and analyzed for DNA contents. The results

indicated that the number of cells was higher in the G2/M phase in N gene transfected

122

cells compared to controls. This suggests that the nucleocapsid protein of arteriviruses

may be associated with cell cycle modulation in arterivirus infected cells. The N proteins

of all members of the family Arteriviridae affected cell cycle in a similar way but their

influence on the cell cycle modulation was different. In the BHK-21 cells expressing the

N protein, PRRSV N and EAV N resulted in more accumulation of cells at G2/M stage

compared to LDV N and SHFV N.

Taken together, it seems that the nucleocapsid protein of Arteriviridae is a

multifunctional protein. It is a nuclear-cytoplasmic protein, and binds to cellular proteins

which are involved in regulation of cell functions, such as transcription and cell cycle. N

protein also affects cell cycle by accumulation of cells at the G2/M stage. These findings

raise the possibility that N protein plays a role in pathogenesis of these viruses, and

further studies are needed to scrutinize its mechanism of function in host cells and

pathogenesis of arteriviruses.

One of the caveats of this research was the lack of an in vivo study for the N

protein nuclear localization and its behavior in primary cells of the natural host. Also, the

cell cycle modification by arteriviruses could have been more deeply investigated.

Therefore, future approaches to deciphering the role of arterivirus N in cell cycle could

be by focusing on each stage of cell cycle. This may be done by synchronizing cells at

G0/G1, S, and G2/M phases followed by virus infection and N protein transfection.

Furthermore, live cell microscopy for monitoring cellular distribution of N protein at

different stages of cell cycle could open another view on its behavior and function in host

cells.

123

In conclusion, the results obtained from the present study shed light on the

subcellular distribution of N protein of arteriviruses. The conserved property of nucleolar

localization of N protein suggests a significant role for this protein in virus pathogenesis.

The finding that N protein is involved in cell cycle modification, supports the assumption

that in addition to its structural role for virus assembly, N protein plays a non-structural

role during virus replication.

124

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