Virus - host cell interactions in echovirus 1 infection

VIRUS - HOST CELL INTERACTIONSIN

ECHOVIRUS 1 INFECTION

Vilja Pietiäinen

University of Helsinki 2005

Helsinki University Biomedical Dissertations No. 63

Virus - host cell interactionsin

echovirus 1 infection

byVilja Pietiäinen

Department of VirologyHaartman Institute

Faculty of Medicineand

Division of BiochemistryDepartment of Biological and Environmental Sciences

Faculty of BiosciencesUniversity of Helsinki

Finland

Helsinki Graduate Schoolin Biotechnology and Molecular Biology

Academic dissertation

To be presented, with the permission of the Faculty of Biosciences of theUniversity of Helsinki, for public criticism in the Small Lecture Hall of the

Haartman Institute on June 10th, 2005, at 12 o'clock noon.

Helsinki 2005

Supervised byProfessor Timo HyypiäDepartment of VirologyUniversity of TurkuandDepartment of VirologyHaartman InstituteUniversity of HelsinkiFinland

Reviewed byProfessor Elina IkonenInstitute of BiomedicineUniversity of HelsinkiFinlandandDocent Maarit SuomalainenDepartment of VirologyHaartman InstituteUniversity of HelsinkiFinland

OpponentProfessor Urs GreberInstitute of ZoologyUniversity of ZürichSwitzerland

ISBN 952-91-7944-8 (paperback)ISBN 952-10-2439-9 (PDF)ISSN 1457-8433http://ethesis.helsinki.fiYliopistopaino, Helsinki 2005

"No matter how hard you try to see, thefuture only reveals itself bit byexasperating bit. The solution to thisanguish is to seize each day with allthe strength, warmth, beauty andloving that is at your command, andyou will make the future happen foryou."

Contents

4

CONTENTS

LIST OF ORIGINAL PUBLICATIONS ..........................................................6

ABBREVIATIONS ..............................................................................................7

ABSTRACT ..........................................................................................................8

TIIVISTELMÄ (FINNISH SUMMARY) .........................................................9

REVIEW OF THE LITERATURE .................................................................10

1. INTRODUCTION TO WORLD OF PICORNAVIRUSES .................................................... 101.1 SHORT HISTORY AND CLASSIFICATION OF VIRUSES .............................................. 101.2 COMMON PROPERTIES OF PICORNAVIRUSES ......................................................... 11

Classification .......................................................................................................... 11Structure.................................................................................................................. 14Viral genome........................................................................................................... 16Replication cycle..................................................................................................... 17

2. PICORNAVIRUS -RECEPTOR INTERACTIONS .............................................................. 202.1 GENERAL PRINCIPLES OF VIRUS-RECEPTOR INTERACTIONS ................................. 202.2 INTEGRINS AS PICORNAVIRUS RECEPTORS ............................................................ 23

Structure and function of integrins ........................................................................ 23αv integrins bind picornaviruses via a viral RGD motif....................................... 25α2β1 integrin as a receptor for EV1...................................................................... 26

2.3 OTHER PICORNAVIRUS RECEPTORS ....................................................................... 28Picornavirus receptors of Ig superfamily bind to the virus canyon...................... 28Other examples of picornavirus receptors............................................................. 31

3. INTERNALIZATION OF PICORNAVIRUSES INTO HOST CELLS ................................... 323.1 CLATHRIN -MEDIATED ENDOCYTOSIS ................................................................... 343.2 LIPID RAFT -MEDIATED ENDOCYTOSIS .................................................................. 363.3 CAVEOLAE -MEDIATED ENDOCYTOSIS .................................................................. 38

4. HOST CELL GENE EXPRESSION DURING PICORNAVIRUS INFECTION ...................... 414.1 CDNA ARRAY STUDIES OF HOST CELL GENE EXPRESSION IN ENTEROVIRUS

INFECTION .................................................................................................................... 41cDNA arrays ........................................................................................................... 42Studies of enterovirus infection with cDNA arrays ............................................... 43

4.2 THE EFFECTS OF EV1 ON HOST CELL GENE EXPRESSION...................................... 46

AIMS OF THE STUDY ....................................................................................47

Contents

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MATERIALS AND METHODS ......................................................................481. VIRUSES (I-IV)......................................................................................................... 482. CELL CULTURES (I-IV) AND TRANSFECTIONS (II, III) ............................................ 483. INFECTIVITY AND BINDING ASSAYS (I-IV).............................................................. 494. SUCROSE GRADIENT SEDIMENTATION (I-III) .......................................................... 515. IMMUNOFLUORESCENCE MICROSCOPY (I-III) ......................................................... 516. REAL-TIME FLUORESCENCE MICROSCOPY (III)....................................................... 527. FLUORESCENCE IN SITU HYBRIDISATION (III)......................................................... 538. PROTEIN SYNTHESIS ASSAY (IV) ............................................................................. 53 9. CDNA ARRAY TECHNIQUE (IV).............................................................................. 53

RESULTS AND DISCUSSION........................................................................54

1. CELL SURFACE INTERACTIONS OF ECHOVIRUS 1 (I-III) ......................................... 541.1 α2β1 INTEGRIN AS EV1 RECEPTOR (I-III) ............................................................ 541.2 The α2I DOMAIN BINDS TO THE EV1 CANYON (I) ................................................ 551.3 THE DIFFERENCES IN INTERACTIONS OF α2β1 INTEGRIN WITH EV1 AND

COLLAGEN (I) ............................................................................................................... 581.4 EV1 BINDING TO α2β1 INTEGRIN MAY TRIGGER INTEGRIN CLUSTERING (I) ....... 591.5 THE UNCOATING OF EV1 IS NOT TRIGGERED BY CELL SURFACE INTERACTIONS

(I-III) ........................................................................................................................... 59

2. THE ENDOCYTOSIS OF ECHOVIRUS 1 INTO CAVEOSOMES (II, III) ......................... 612.1 EV1 DOES NOT UTILIZE CLATHRIN-MEDIATED ENDOCYTOSIS (II, III) ................. 612.2 EV1 MAY UTILIZE BOTH CELL SURFACE CAVEOLAE AND AN ALTERNATIVE

PATHWAY TO ENTER HOST CELLS (II, III).................................................................... 622.3 EV1 IS INTERNALIZED INTO CAVEOSOMES (II, III) ............................................... 652.4 EV1 REMAINS IN CAVEOSOMES PRIOR TO REPLICATION (III)............................... 67

3. ECHOVIRUS 1 INFECTION RESULTS IN ALTERATIONS OF HOST CELL GENE

EXPRESSION (IV) ............................................................................................................. 693.1 THE UPREGULATION OF IMMEDIATE EARLY GENES .............................................. 703.2 THE UPREGULATION OF IRES-CONTAINING CELLULAR GENES............................ 713.3 OTHER UPREGULATED GENES................................................................................ 71

CONCLUSIONS ................................................................................................74

ACKNOWLEDGEMENTS ..............................................................................77

REFERENCES...................................................................................................79

ORIGINAL PUBLICATIONS .........................................................................93

List of Original Publications

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LIST OF ORIGINAL PUBLICATIONS

This thesis is based on the following original publications, which are referredto in the text by their Roman numerals (I-IV).

I Xing L., Huhtala M., Pietiäinen V., Käpylä J., Vuorinen K., Marjomäki V.,Heino J., Johnson M.S., Hyypiä T., and Cheng R.H. 2004. Structural andfunctional analysis of integrin α2I domain interaction with echovirus 1.Journal of Biological Chemistry 279:11632-11638.

II Marjomäki, V., Pietiäinen, V., Matilainen, H., Upla, P., Ivaska, J.,Nissinen, L., Reunanen, H., Huttunen, P., Hyypiä, T., and Heino, J. 2002.Internalization of echovirus 1 in caveolae. Journal of Virology 76:1856-1865.

III Pietiäinen V., Marjomäki V., Upla P., Pelkmans L., Helenius A., andHyypiä T. 2004. Echovirus 1 endocytosis into caveosomes requires lipid rafts,dynamin II, and signaling events. Molecular Biology of the Cell 11:4911-25.

IV Pietiäinen, V., Huttunen P., and Hyypiä, T. 2000. Effects of echovirus 1infection on cellular gene expression. Virology 276: 243-250.

The original papers have been reprinted with the kind permission of thecopyright holders.

Abbreviations

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ABBREVIATIONS

AF Alexa Fluor fluorescent dyeα2I I domain of the integrin α2 subunitATCC American Type Culture Collectionβ2m β2 microglobulinCAR coxsackievirus and adenovirus receptorCAV coxsackie A virusCBV coxsackie B viruscDNA complementary DNACPE cytopathic effectCTX cholera toxinCV-1 African green monkey kidney cell lineDAF decay-accelerating factorEE early endosomeEM electron microscopyER endoplasmic reticulumERK extracellular signal-regulated kinaseEV echovirusFISH fluorescence in situ hybridizationFMDV foot-and-mouth disease virusGFP green fluorescent proteinGMK African green monkey kidney cell lineGST glutathione-S-transferaseHOS human osteosarcoma cell lineHPEV human parechovirusHRV human rhinovirusICAM-1 intercellular adhesion molecule-1IE immediate early (genes)IF immunofluorescenceIg immunoglobulinIRES internal ribosome entry siteLE late endosomeMAb monoclonal antibodyMAPK mitogen-activated protein kinaseMEM minimal essential mediumMIDAS metal-ion-dependent adhesion sitemRNA messenger RNAPAGE polyacrylamide gel electrophoresisPBS phosphate-buffered salinep.i. post infection (indicates time after 1-h incubation of EV1 at 0-4°C)PV poliovirusPVR poliovirus receptorSAOS human osteosarcoma cell lineSDS sodium dodecyl sulphateSV40 simian virus 40VLDL-R very-low-density lipoprotein receptorVP viral proteinVWA von Willebrand A domainsWT wild type

Abstract

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ABSTRACT

The family Picornaviridae consists of many clinically and economicallysignificant pathogens of humans (polioviruses, hepatitis A virus,rhinoviruses) and live-stock (foot-and-mouth disease viruses). Studies onthese small, non-enveloped animal viruses have made a great impact on thedevelopment of modern virology. The first steps of picornavirus infectioninclude binding to cell surface receptor, entry and uncoating of virus. Theuncoating leads to the release of the positive-stranded viral genome, whichcan directly act as a messenger RNA in the translation. The viral replicationoccurs in the cytoplasm, where newly synthesized viral capsid proteins andgenome are assembled. The progeny virus particles are, in most cases,released by lysis of the host cell.

This study focuses on host cell interactions of echovirus 1 (EV1), a memberof the enterovirus genus of Picornaviridae. On the cell surface, EV1 binds tothe α2I domain of α2β1 integrin, a collagen receptor. In the first phase of thethesis, the virus-integrin interactions were investigated by cryo-electronmicroscopy remodelling in a collaborative study. The binding site of α2Idomain was defined as the top of the canyon structure in the EV1 capsid. Theresults indicated that there were significant differences in the bindingmechanisms of EV1 and collagen to the integrin. Binding of EV1 to the α2Idomain or on the cell surface did not trigger disassembly of viral capsid andrelease of the viral RNA. The results gave new insights into picornavirus-receptor interactions and into integrin-ligand interactions in general.

In the second and the third phases of the study it was found that EV1 isinternalized into host cells via the cell surface caveolae or by an alternative,unknown pathway. Caveolae-mediated endocytosis is also important forcellular functions, however, the detailed mechanisms of the pathway are notyet thoroughly understood. Both entry pathways of EV1 are dependent on thepresence of α2β1 integrin and they guide the virus into intracellular vesicles,caveosomes. The real-time live microscopy revealed the rapid uptake offluorescently labelled EV1 into these structures. Interestingly, the virusremained in caveosomes prior to the initiation of viral replication. EV1endocytosis represents a new model for picornavirus entry and for cellularendocytic events.

Viruses can have dramatic effects on host cell gene expression. In the fourthphase, such effects during EV1 infection were investigated using cDNA arrayanalysis. Changes in host cell gene expression included increased synthesis ofimmediate early response genes and genes involved, e.g., in stress responsepathways. EV1 caused also a partial shut-off of host cell protein synthesis, amechanism to increase the viral replication efficiency. The results implicatedthat EV1 infection has multiple effects on the host cell that might beconsequences of both host cell defence and viral replication.

Finnish Summary

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TIIVISTELMÄ (FINNISH SUMMARY)

Virusten on sitouduttava solunpinnan vastaanottajamolekyyliin, reseptoriin,aloittaakseen lisääntymiskierron isäntäsolussa. Tämän jälkeen virus meneeisäntäsolun sisään esimerkiksi endosytoosin avulla. Näin virus pääseemäärättyyn solunsisäiseen paikkaan, jossa se lisääntyy tuottaakseen uusiaviruspartikkeleita. Tuotetut viruspartikkelit vapautuvat solusta aloittaakseentaas infektiokierron uusissa kohdesoluissa.Echovirus 1 (EV1) on pieni (30 nm) vaipaton pikornavirus, jolla onpositiivijuosteinen RNA-perintöaines. Pikornavirusheimoon kuuluvat myössellaiset merkittävät ihmisten ja eläinten taudinaiheuttajat kuten poliovirukset,rinovirukset, hepatiitti A -virus ja karjan suu- ja sorkkatautivirus. Tässäväitöskirjatyössä on tarkasteltu EV1:n solukierron eri vaiheita: viruksensitoutumista reseptoriin, endosytoosia, sekä virusinfektion vaikutuksia solungeenien ilmentymiseen.EV1 sitoutuu solun pinnalla α 2β1-integriiniin, joka toimii normaalistikollageenireseptorina. Tarkka sitoutumispaikka integriinissä sijoittuu α2-alayksikön I-domeeniin (α2I). Tämän väitöskirjan ensimmäinen osatyö olikansainvälinen yhteistyöhanke, jossa määritettiin sekä EV1-α2I domeeninvälisen vuorovaikutuksen biokemiallisia ominaisuuksia että α2I-domeeninsitoutumispaikka viruksessa kryo-elektronimikroskopian avulla. Tulostenperusteella α2I-domeeni sitoutuu kanjoniksi kutsuttuun syvänteeseen viruksenkapsidissa.Toisessa ja kolmannessa osatyössä tutkittiin EV1:n tunkeutumistaisäntäsoluun, mm. elävissä soluissa fluoresoivasti leimatun viruksen avulla.Integriiniin sitoutuneen EV1:n osoitettiin menevän kaveoli -välitteisenendosytoosin tai tuntemattoman vaihtoehtoisen reitin kautta solun sisäisiinkaveosomirakenteisiin. Virus pysyi kaveosomeissa ennen RNA-genominmonistumista solulimassa. Tähän mennessä EV1 on ainoa pikornavirus, jonkaon näytetty käyttävän kaveolireittiä, ja siten työ tarjoaa uuden mallinpikornaviruksen soluunmenolle ja antaa myös lisätietoa solunkuljetusmekanismeista.Virukset muokkaavat usein solun perustoimintoja ja valjastavat solunproteiineja omaan käyttöönsä. Neljännessä osatyössä tutkittiin EV1:nlisääntymiskierron vaikutuksia isäntäsolun geenien ilmentymiseen cDNA-siruteknikkaa (cDNA array) käyttäen. Viruksen lisääntyminen isäntäsolussavaikutti eniten isäntäsolun geenien ilmentymisen muutoksiin. Muutoksianähtiin esimerkiksi ohjattuun solukuolemaan, solun stressitilanteisiin ja solunkasvun säätelyyn liittyvien geenien ilmentymisessä. EV1 myös esti osittainisäntäsolun proteiinivalmistuksen parantaakseen lisääntymistehokkuuttaan.Havaitut muutokset isäntäsolussa infektion aikana voivat johtua sekä itseinfektiosta että solun omista torjuntamekanismeista.

Review of the Literature

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REVIEW OF THE LITERATURE

1. Introduction to world of picornaviruses

1.1 Short history and classification of viruses

Viruses are small, infectious agents that can multiply within the cells of

humans, animals, plants, and in bacteria. The first evidence of viruses was

obtained in the 1890s by Dimitrie Ivanowski and Martinus Beijerinck. They

reported that a pathogenic agent associated with tobacco mosaic disease of

plants passed through filters that trapped all known bacteria (reviewed in

Lederberg, 2000). During the same decade, Friedrich Loeffler and Paul

Frosch found that the agent causing foot-and-mouth disease in livestock was

also filterable (reviewed in Mahy, 2005). In 1901, Walter Reed and James

Carrol discovered the viral origin for the serious human disease, yellow fever

(Lederberg, 2000). The virological background of poliomyelitis was

established by Landsteiner and Popper in 1908 when they succeeded to

transfer poliomyelitis from human samples to monkeys in a form of filterable

agent (reviewed in Flint et al., 2000). These investigations initiated

identification of a large number of viruses infecting many organisms.

During the last two centuries, the spectrum of circulating viruses has changed

when the hygiene has improved and potent vaccines have been developed.

However, viruses are still causing many millions of deaths every year because

of several reasons. Viruses evolve all the time, new and emerging viral

epidemics take place and increased travelling enables efficient transmission of

viruses from one continent to another (Lee and Henderson, 2001). On the

other hand, some lethal viral diseases have been completely eradicated from

the world, e.g. the smallpox that caused over 300 million cases in the 20th

century (Mahalingam et al., 2004). Polio eradication is under way (Minor,

2002) and the next target will be measles as assigned by the World Health

Organization (de Quadros, 2004). However, influenza virus is still causing

severe epidemics almost every year (Kilbourne, 2004) and human


11

immunodeficiency virus, HIV, spreads especially in developing countries

(Steinbrook, 2004).

The extensive epidemics have not only affected humans, but they have caused

serious economical damage also to agriculture by targeting both animals and

cultivated plants. Thus, viruses are mostly regarded as uninvited and

opportunistic guests, which struggle with their host in an effort to replicate

and spread their genetic material. On the other hand, the simplicity of viruses

has made them instruments for gene therapy and for studies on immunology

and cellular and molecular biology (Pelkmans and Helenius, 2003). For

example, discovery that propagation of poliovirus is possible in cell culture

significantly encouraged researchers towards cell biological studies of animal

viruses in 1950s and made development of efficient vaccines possible.

Viruses are classified, for example, based on 1) nature of viral nucleic acid

and replication strategy, 2) presence or absence of an envelope and 3) virion

and nucleocapsid morphology (Condit, 2001). The size of viruses varies from

15 to 300 nm. The viral capsid encloses 3 to 300 kilobases long, a circular or

linear DNA or RNA genome that exists in one piece or in a segmented form.

If the nucleic acid is single-stranded, it can be either positive (+) (same sense

as mRNA) or negative-stranded (-) (complementary to mRNA). Viral capsid

is composed of copies of one or more structural proteins and it displays

icosahedral (spherical viruses), helical (rod-shaped viruses) or more complex

symmetry. In addition, some viruses have a lipid envelope, derived from the

cell membrane through which the viral capsid has budded.

1.2 Common properties of picornaviruses

ClassificationThis thesis concentrates on the cellular interactions of echovirus 1 (EV1), a

member of family Picornaviridae. Picornaviruses are small (30 nm), non-

enveloped animal viruses. They carry a positive-stranded RNA genome that

can directly act as an mRNA when released into a host cell. Picornaviruses


12

are divided into nine genera: aphtho-, cardio-, entero-, erbo-, hepato-, kobu-,

parecho-, rhino-, and teschoviruses (King, 2000) (Table 1.) Each genus is

further divided into species, which consist of different virus serotypes with

distinct antigenic determinants. Previously, picornaviruses were classified

mainly according to their pathogenesis in laboratory animals (Hyypiä et al.,

1997) but a current classification is based on genetic information of the virus.

Picornaviruses cause a great variety of diseases in humans and other animals,

varying from hepatitis (hepatitis A virus; HAV), poliomyelitis (polioviruses;

PVs) and common cold (human rhinoviruses; HRVs) to foot-and-mouth-

disease of cattle (foot-and-mouth-disease viruses; FMDVs) (Table 1). In

clinical terms, enteroviruses, to which EV1 belongs, are probably the most

significant members of the picornavirus family. Human enteroviruses (HEV)

contain five different species: polioviruses and HEVs A to D. In addition, the

enterovirus genus contains three species of viruses of other animals: bovine

enterovirus, porcine enterovirus A and porcine enterovirus B (King, 2000;

Stanway et al., 2002).

Enteroviruses cause a wide spectrum of illnesses among new-borns, children

and adults, including respiratory infections, poliomyelitis, meningitis,

encephalitis, myocarditis and conjunctivitis. In addition, increasing data

suggest that enteroviruses may have a role in the development of type 1

diabetes (Hyöty and Taylor, 2002). However, the majority of infections are

asymptomatic (Grist et al., 1978). The faecal-oral route is the main way of

enterovirus transmission. Primary replication of enteroviruses takes place in

respiratory and gastrointestinal tissues. Viremia is caused when the viruses

circulate in the blood stream and can reach target organs, such as liver, heart

and central nervous system (Pallansch and Roos, 2001). Polioviruses (PVs),

the best-studied species among enteroviruses, cause poliomyelitis, a serious

disease that has affected humans for thousands of years. Fortunately, highly

efficient vaccines against PVs have been developed and the worldwide effort

to eradicate polio is likely to reach its goal during the next few years. In June

2002, the WHO European region was certified polio-free, as are the regions of

the Americas and the Western Pacific (Minor, 2002).


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Table 1. Classification of picornaviruses & examples of clinical diseases associatedwith picornavirus infections (Grist et al., 1978; King, 2000; Stanway et al., 2002).

Genus Species (No of serotypes) Clinical DiseasesEnterovirus

Poliovirus (3) (PVs)- Human polioviruses 1-3

Poliomyelitis

Human enterovirus A (12) (HEV-A)- Human coxsackieviruses A2-A8, A10-A14, A16- Human enterovirus 71

Meningitis, paralysis,myocarditis, rash

Human enterovirus B (37) (HEV-B)- Human coxsackieviruses B1-B6- Human coxsackievirus A9- Human echoviruses 1-7, 9, 11-21, 24-27, 29-33- Human enteroviruses 69, 73

Meningitis, paralysis,myocarditis,gastroenteritis

Human enterovirus C (11) (HEV-C)- Human coxsackieviruses A1, A11, A13, A15,A17-A22, A24

Respiratory infections,conjunctivitis

Human enterovirus D (2) (HEV-D)- Human enteroviruses 68, 70

conjunctivitis

Bovine enterovirus (2)Porcine enterovirus A (1)Porcine enterovirus B (2)

RhinovirusHuman Rhinovirus A (74) (HRVs) Common coldHuman Rhinovirus B (25) Common cold

CardiovirusesEncephalomyocarditis virus (1) CarditisTheilovirus (2)

AphthovirusFoot-and-mouth disease virus (7) (FMDVs) Foot- and- mouth diseaseEquine rhinitis A virus (1)

HepatovirusesHepatitis A virus (1) (HAV) Liver disease

ParechovirusesHuman parechovirus (2) (HPEVs) Respiratory infections,

gastroenteritisLjungan virus (2?)

ErbovirusEquine rhinitis B virus (2)

KobuvirusAichi virus (1) Gastroenteritis

TeschovirusPorcine teschovirus (10)

All echovirus (enteric cytopathogenic human orphan) serotypes, including

EV1, belong to HEV-B species (King, 2000). Echoviruses were initially

distinguished from coxsackieviruses by their inability to replicate and cause

disease in newborn mice (Pallansch and Roos, 2001). Echovirus infections are

most often subclinical, although the viruses may cause clinical diseases such

as aseptic meningitis, muscle weakness and paralysis, exanthemas,

pericarditis, myocarditis, common cold, uveitis, conjunctivitis, infantile

diarrhoea, and acute febrile respiratory illness (Grist et al., 1978). The


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outbreaks are most common among neonates and infants (Pallansch and Roos,

2001). Recently, a mouse-model expressing the EV1 receptor, α2β1 integrin,

was developed, enabling establishment of a pathogenetic model for EV1

(Hughes et al., 2003). Intracerebral inoculation of new-born transgenic mice

with EV1 led to paralysis, whereas adolescent mice did not display

neuropathology or paralytic disease but developed myocarditis.

StructurePicornaviral capsid is an icosahedral, spherical particle with a diameter of

about 30 nm (Fig. 1A, B). The atomic structure of EV1 was recently

determined by cryo-crystallography (Filman et al., 1998). The other atomic

structures of picornaviruses that have been elucidated include CAV9 (Hendry

et al., 1999), CAV21 (Xiao et al., 2001), CBV3 (Muckelbauer et al., 1995),

EV11 (Stuart et al., 2002b), FMDV (Acharya et al., 1989), HRV2 (Verdaguer

et al., 2000), HRV14 (Rossmann et al., 1985), PV1 (Hogle et al., 1985) and

PV3 (Filman et al., 1989). The structure of PVs is best characterized among

enteroviruses and shows their common structural properties.

Similarly to other picornaviruses, the capsid of EV1 is composed of 60

heteromeric structural units, protomers (Filman et al., 1998). A protomer

consists of single copies of each of four capsid proteins VP1-VP4 (Fig. 1B).

Five protomers assemble into a pentamer, and twelve pentamers form a viral

capsid. VP1, VP2 and VP3 decorate the outer surface of the capsid while VP4

is buried inside and represents the detached amino-terminal extension of VP2.

Even though the primary structures of VP1-VP3 are different, each of capsid

proteins contains two α-helices and a similar, eight-stranded, anti-parallel β-

barrel core (Hogle et al., 1985) (Fig. 1C). The loops connecting β-strands are

highly diverse, particularly at the top of the β-barrel domain. The loops and

the C-terminus create the surface features, antigenic properties and receptor

recognition characteristics of the virion.

The five-fold axis is formed of β-barrels from five copies of VP1, whereas the

three-fold axis is surrounded by VP2 and VP3 (Hogle et al., 1985) (Fig. 1B).

The plateau of five-fold axis is encircled by a depression. In echoviruses,

related enteroviruses, and in a major group of rhinoviruses these depressions


15

are joined to form a canyon, which is the site of receptor attachment for many

picornaviruses (Rossmann et al., 2002) (Fig. 1B). The "canyon hypothesis"

proposed that the receptor binds to the conserved sequences on the bottom of

the canyon and thus escapes the neutralizing host antibodies that are too large

to reach the receptor-binding site (Rossmann, 1994). Later, this hypothesis

was challenged as receptor and antibody binding sites partially overlap on

capsid surface (Rossmann et al., 2002).

Figure 1. Structure of picornaviruses. A) Electron micrograph of negatively stainedEV1 shows small, spherical viral particles. Scale bar 100 nm. B) The picornavirus(EV1) capsid consists of 60 protomers. VP1, VP2 and VP3 of one protomer areindicated in the virus capsid. VP4 is buried inside the capsid. Three-fold (3) and five-fold (5) symmetry axes and the canyon are indicated by arrows. (The model of EV1capsid was adapted from http://mmtsb.scripps.edu/viper/; copyright © 1998-2004 byTSRI) C) Capsid proteins VP1, VP2, and VP3 share a common core structure of eightβ-strands, connected by loops (Modified from Hogle, 2002).

The N-terminal residue of VP4 of all studied enteroviruses, including EV1, is

covalently bonded to a myristic acid group (Chow et al., 1987). This fatty

acid may play a role in the capsid assembly and entry events. Moreover, in

most enteroviruses and in a major group of rhinoviruses, the hydrophobic

bottom of the canyon contains a pocket-factor of cellular origin (Filman et al.,


16

1989). The pocket factor (=lipid) stabilizes the capsid and it is released before

uncoating (Racaniello, 2001). Thus it may prevent premature uncoating and

RNA release and also ensure that viruses are carried from the cell in an intact

form.

Viral genome

The size of the single-stranded (+)RNA of picornaviruses varies from 7.2

(HRV14) to 8.5 (FMDV) kilobases. The genome contains a single open

reading frame (Fig. 2).

Figure 2. Genomic structure of enteroviruses. The P1 region (white) is processed tostructural proteins VP1-VP4. The P2 and P3 regions (gray) are processed tononstructural proteins. The principal functions of viral proteins during infection cycleare presented in boxes. (Modified from Bedard and Semler, 2004).

The 3' and 5' ends of the viral RNA contain untranslated regions (UTRs). The

3' end of picornaviruses carries a PolyA -sequence (Yogo and Wimmer,

1972), and a secondary structure (pseudoknot), used for initiating the

synthesis of negative strand RNA (Jacobson et al., 1993). The 5' end of the

viral RNA is not capped like cellular mRNAs, but instead it contains a

covalently attached viral protein 3B, also called VPg (Nomoto et al., 1977).

An internal ribosome entry site (IRES), which is required for cap-independent

translation(Pelletier and Sonenberg, 1988), and the clover-leaf structure,

which is involved in RNA replication (Andino et al., 1990), are also present at

the 5' end.


17

The enterovirus genome is translated to one polyprotein, which is further

cleaved to precursor proteins P1, P2 and P3. P1 precursor protein is cleaved

into the structural proteins VP1-VP4. The nonstructural proteins, which are

required for proteolytic cleavages, for viral translation and for RNA

replication, are formed from precursors P2 (2A, 2B, and 2C) and P3 (3A, 3B,

3C and 3D) (Racaniello, 2001). The 3D is an RNA-dependent RNA-

polymerase, which specifically copies viral RNA in the presence of VPg

primer (Baltimore et al., 1963; Nomoto et al., 1977). The 2A acts as a

protease in enteroviruses and rhinoviruses and the 3C in all picornaviruses

(Hanecak et al., 1982; Toyoda et al., 1986).

Replication cycleAttachment (1) and entry (2). Viral infection starts by interaction of virus

with its cell surface receptor or multiple receptors (Fig. 3.). In picornaviruses,

the receptor interactions may result in conformational changes essential for

viral entry and RNA release (=uncoating) (Rossmann et al., 2002). Uncoating

may also be triggered by other factors, such as acidification. The receptor-

triggered uncoating of PVs may be followed by penetration of viral RNA into

the cytoplasm through a pore within the plasma membrane (Hogle, 2002).

Alternatively, picornaviruses can be endocytosed into the host cell prior to the

uncoating. After viral genomic RNA is released into the cytoplasm, the

genomic VPg is cleaved. The (+)RNA acts directly as mRNA for synthesis of

the viral polyprotein precursor.

Translation (3). Since cellular proteins cannot copy picornaviral RNA, it

must first be translated in order to produce viral proteins required for

replication of the viral genome. Picornaviruses inhibit the cellular protein

synthesis by cleaving the cellular components, which are essential for cap-

dependent cellular translation (Etchison et al., 1982; Gradi et al., 1998).

However, the cap-independent translation of viral proteins is allowed due to

the presence of IRES in the enterovirus genome (Dorner et al., 1984; Pelletier

and Sonenberg, 1988). In addition to canonical initiation factors, some

noncanonical factors, such as poly(rC) binding protein, are required for

initiation of viral translation (Racaniello, 2001). The instant cleavage of the


18

Figure 3. Life cycle of picornaviruses: 1) Attachment, 2) Entry 3) Translation, 4)

Replication, 5) Assembly and 6) Release. Each number is referred to in the text.

(Modified from Flint et al., 2000).


19

translated polyprotein is performed by virus-encoded proteinases (Kitamura et

al., 1981; Semler et al., 1981). The non-structural proteins (except 2A), which

are cleaved from precursors P2 and P3, participate in viral RNA replication.

Replication (4). Picornaviruses, like other RNA viruses, have membrane-

associated replication complexes (Egger et al., 2002). With picornaviruses,

the rosette-like complexes consist of replicating viral RNA, viral and cellular

proteins, and tubulated, virus-induced membranous vesicles. The replication

is initiated by a complex of viral and host cell proteins bound to 5' cloverleaf

of viral RNA (Andino et al., 1999). Viral 3Dpol copies genomic (+)RNA into

complementary (-)RNAs, which carry VPg at their 5' ends and serve as

templates for newly synthesized genomic (+)RNA. Simultaneously, VPg is

removed from some of the newly synthetized (+)RNAs that are further

translated for efficient production of viral proteins.

Assembly (5) and the release (6). A precursor protein P1 is further cleaved

into coat proteins VP0 (a precursor of VP4 and VP2 in most of the

picornaviruses), VP3 and VP1. Protomers, carrying one copy of each coat

protein, associate with genomic RNA containing VPg to form progeny viruses

(Racaniello, 2001). Finally, VP0 is cleaved into VP4 and VP2 for production

of infectious viral particles. Picornaviruses, like most nonenveloped viruses,

are usually released from cells by cell lysis.

The replication cycle of picornaviruses takes approximately 6 to 12 hours in

cells. About 50 000 new viral particles are produced during each cycle in one

cultured cell but only 0.1-2% of them are infectious (Racaniello, 2001). This

may be due to the lethal mutations in the viral genome and/or other defects of

the infectious cycle.


20

2. Picornavirus-receptor interactions

2.1 General principles of virus-receptor interactions

In order to enter into host cells and initiate infection, viruses must attach to

the specific receptor(s) on the cell surface. These receptors can often have

important functions in cell adhesion, cell-cell interactions, signalling and

defence mechanisms. The binding of virus to a receptor can elicit changes in

receptor conformation. These alterations may bring about signalling events

that regulate both the viral entry process and the cellular response to the

infection. On the other hand, conformational changes in virus particles,

triggered by receptor binding, can also facilitate virus entry and uncoating.

Among the best characterized virus-receptor interactions are those of HIV

with CD4 molecule and chemokine receptors (reviewed in Smith and

Helenius, 2004). Binding of HIV to CD4 molecule on the cell surface leads to

conformational changes in virus structure. These changes allow the further

interactions of HIV with chemokine receptors that in turn promote the fusion

of viral envelope with the plasma membrane and subsequent release of viral

core into the cytoplasm.

Picornaviruses can interact with a great variety of cell surface molecules,

including members of immunoglobulin superfamily (IgSF) and integrins

(Evans and Almond, 1998)(Table 2 and Fig. 4). Many picornaviruses share

cellular receptors, for instance, decay accelerating factor (DAF) functions as a

receptor for several enteroviruses, and αv-integrins are utilized by FMDVs,

human parechovirus 1 (HPEV1) and coxsackievirus (CAV) 9. Furthermore,

picornaviruses and other viruses can utilize the same receptors; for example,

both CBVs and adenoviruses bind to coxsackievirus-adenovirus receptor,

CAR (Tomko et al., 1997). Often, one receptor is required for binding and

another for uncoating and entry (Rossmann et al., 2002): e.g. CAV21 binds

primarily to DAF but enters the cells via an intercellular adhesion molecule-1

(ICAM-1) (Shafren et al., 1997). Alternative receptor(s) may also be utilized

depending on cell type and cell polarization. For example, CBV3 can infect

the apical surface of polarized epithelium which expresses its secondary


21

receptor DAF (Shafren et al., 1995), even though a primary receptor, CAR

(Bergelson et al., 1997; Tomko et al., 1997), is hidden within intercellular

junctions (Shieh and Bergelson, 2002). However, for some picornaviruses,

like PVs (Mendelsohn et al., 1989) and a major group of HRVs (Greve et al.,

1989), one receptor is enough to ensure attachment, uncoating and entry.

Table 2. The examples of cell surface receptors for picornaviruses. (Modified fromRacaniello, 2001).

RECEPTOR(S) VIRUSIgSF-like- Poliovirus receptor; CD155, PVR Polioviruses 1,2,3- Intercellular adhesion molecule -1; ICAM-1 Coxsackieviruses A13, A18, A21

Major group of rhinoviruses- Coxsackievirus-adenovirus receptor; CAR Coxsackieviruses B1-B6- HAV cellular receptor 1; HAVcr-1 Hepatitis A virus

SRC-like- Decay accelerating factor; CD55; DAF Coxsackievirus A21*

Echoviruses 3, 6, 7, 11-13, 20, 21, 24, 29, 30Enterovirus 70Coxsackieviruses B1, B3, B5*

Integrins− α2β1 integrin Echovirus 1− αvβ3 and αvβ6 integrin Coxsackievirus A9− αvβ1 and αvβ3 integrin Parechovirus 1− αvβ1, αvβ3, αvβ6 and α5β1 integrin Foot-and-mouth disease virus (field isolates)

Signalling receptors- Low-density lipoprotein receptor; LDL-R Minor group of rhinoviruses

Carbohydrates- Sialic acid Rhinovirus 87

Enterovirus 70*

Glycosaminoglycans- Heparan sulphate Foot-and-mouth disease virus (culture

adapted)Echovirus 6 and certain other serotypes*

Others− β2 microglobulin (β2m) Certain echovirus serotypes*,

Coxsackievirus A9**) the virus uses the receptor as a "secondary" receptor. SRC= short consensus repeat

It is important to keep in mind that the reported virus-receptor interactions are

usually based on cell culture studies of laboratory strains of viruses, which

have adapted to certain receptors and may not be similar to the clinically

circulating viruses. For example, the field isolates of FMDVs bind to integrins


22

while laboratory strains of viruses use heparan sulphate as a receptor (Jackson

et al., 1996; Neff et al., 1998).

Figure 4. Different types of picornavirus receptors. SRC=short consensus repeat,GPI=glycosylphosphatidyl inositol anchor (Modified from Evans and Almond, 1998).

Binding of natural ligands to cell surface molecules is known to induce

conformational changes and, sometimes, clustering of receptors, which may

lead to a variety of signalling events (Greber, 2002). Similarly, virus-receptor

interactions can trigger signalling events that influence virus entry,

cytopathogenecity and the immune response. So far, signal transduction

following the picornavirus-receptor interactions has not been extensively

studied. Interestingly, recent findings suggest that the interactions of EV1

with its receptor, α2β1 integrin, can trigger a cascade of signalling events,

including protein kinase Cα activation (Upla et al., 2004). These signalling

events are required for virus internalization into the host cell (Pietiäinen et al.,

2004; Upla et al., 2004). This is somewhat similar to the signalling events

observed during adenovirus (Ad-2 and Ad-5) interaction with αv-integrins.

The adenovirus-integrin interaction results in an activation of several

signalling pathways and dynamic changes in the actin cytoskeleton, events

which mediate internalization and endocytic trafficking of the virus (Greber,

2002; Goosney and Nemerow, 2003).


23

Although virus-receptor interactions are required for initiation of infection

and may promote the subsequent steps of the viral life cycle, the viral

pathogenesis is not determined only by receptor recognition (Schneider-

Schaulies, 2000). In addition, intracellular factors, the velocity of virus

replication, cytopathogenecity, the spread of infection within and between

organs and the host immune response have an influence on the development

of disease.

2.2 Integrins as picornavirus receptors

Structure and function of integrinsIntegrins are heterodimers of two covalently associated subunits, α and

β (Ruoslahti and Pierschbacher, 1987; Takagi and Springer, 2002) (Fig. 5).

Currently, 24 combinations of 18 integrin α and eight integrin β subunits are

known (Hynes, 2002). The subunits contain a large extracellular domain of

≥940 (α) and ≥640 (β) residues, a single transmembrane domain and a short,

C-terminal, cytoplasmic tail (Fig. 5).

Figure 5. The structure of integrin, which contains the I domain in the α subunit.(Modified from Humphries, 2002; Hynes, 2002).


24

About half of vertebrate integrin α subunits contain an I ("inserted") domain

(Whittaker and Hynes, 2002). I domain is a member of a family of von

Willebrand A domains (VWA). The members of the VWA family share a

common structure of Rossman folds, which consist of a β-sheet sandwiched

between multiple α helices (Lee et al., 1995) (Fig. 6). I domain interacts

with the β-propeller, which is formed of seven similar structural units (∼ 60

amino acids each) (Springer, 1997). The leg the of α-subunit consists of a

thigh domain and two calf domains (Hynes, 2002). The head of all β-subunits

contains an I-like domain, which shares a common structure with αI domains

(Lee et al., 1995) (Fig. 5). The β I-like domain interacts with α -subunit,

forming an interface for a ligand binding. The leg of β-subunit has a hybrid

domain, plexin-semaphorin-integrin (PSI) domain, four cystein-rich repeats

(I-EGF; epidermal growth-factor domains) and a novel cystatin-like fold (β-

tail domain) (Hynes, 2002). A metal-ion binding site (MIDAS), essential for

ligand binding (Michishita et al., 1993), is present in both the αI domain and

β I-like domain.

Integrins are cell-adhesion receptors, crucial for cell invasion, migration and

survival (Hood and Cheresh, 2002). Therefore, they are involved in

developmental processes, immune response, chronic inflammation and

invasion of cancer (Hynes, 2002). Integrins bind many ligands, including a

large number of extracellular matrix proteins (e.g. collagens, fibronectins,

vitronectin, laminins, von Willebrand factor and thrombospondins), counter-

receptors (ICAMs and generally members of the IgSF) and plasma proteins

(Hynes, 1992). Numerous pathogens, including adenoviruses (Wickham et al.,

1993), cytomegaloviruses (Feire et al., 2004), picornaviruses (Bergelson et

al., 1992; Roivainen et al., 1994; Berinstein et al., 1995) and rotaviruses

(Guerrero et al., 2000; Ciarlet et al., 2002) utilise integrins as cell surface

receptors. Many but not all integrin ligands contain an arginine-glycine-

aspartatic acid (RGD) tripeptide that specifically binds to certain integrins,

such as α5β1, αVβ3, αVβ5, αVβ6 and αVβ8 (Ruoslahti and Pierschbacher,

1987).


25

Integrin signalling and activation are both mediated by large conformational

changes that are propagated from the integrin headpiece to the cytoplasmic

domains and vice versa (Hynes, 2002). Thus, integrins can signal in both

directions, outside-in and inside-out. Binding to the matrix induces

association of integrins with the actin cytoskeleton and activates biochemical

signals inside the cell. Conversely, intracellular signals can promote the

binding of integrins to matrix ligands. This inside-out signalling may trigger

transformation of integrins from a closed and inactive "low affinity"

conformation to an open and active "high affinity" state (Takagi and Springer,

2002). The high affinity conformation of integrins is required for binding of

some ligands, e.g. collagen (Emsley et al., 2000). The ligated integrins can

cluster by oligomerization of their transmembrane domains, which results in

the activation of cellular signalling cascades (Qin et al., 2004).

αv integrins bind picornaviruses via a viral RGD motifPicornaviruses CAV9 (Chang et al., 1989), HPEV1 (Hyypiä et al., 1992) and

FMDVs ( F o x et al., 1989) contain an RGD motif (Ruoslahti and

Pierschbacher, 1987), which often offers a binding site for integrins. The

RGD sequence is present in the capsid protein VP1 on the surface of viruses

(Fox et al., 1989; Chang et al., 1992). In contrast to many other picornavirus

receptors, the integrins interacting with RGD-containing picornaviruses do

not appear to bind into the virus canyon.

CAV9 was first shown to utilize the αvβ3 integrin as a receptor (Roivainen et

al., 1991; Roivainen et al., 1994), but, according to later reports, it can also

bind to other α v-integrins, such as αvβ6 (Williams et al., 2004). The

interaction of CAV9 with αvβ6 is RGD-dependent (Williams et al., 2004).

However, RGD is not essential for CAV9 infectivity, as the virus can

efficiently bypass the RGD-dependent entry (Roivainen et al., 1991; Hughes

et al., 1995; Roivainen et al., 1996). This suggests that CAV9 could also use

other receptors for cell entry. Indeed, MHC class I protein as well as two

MHC class I -associated proteins, β2 microglobulin and GRP78 (a member of

heat shock protein-70 family of stress proteins) may be involved in the entry

process of CAV9 (Triantafilou et al., 1999; Triantafilou et al., 2002).


26

HPEV1 was found to compete with CAV9 for the cell surface binding,

therefore leading to the presumption that these viruses may share a common

receptor (Roivainen et al., 1994). Indeed, HPEV1 also utilizes αv-integrins as

receptors (Stanway et al., 1994), e.g. αvβ1 (Pulli et al., 1997) and αvβ3

(Stanway et al., 1994; Joki-Korpela et al., 2001). In contrast to CAV9, the

RGD motif has been shown to be critical for HPEV1 viability (Boonyakiat et

al., 2001). Interestingly, αvβ3 integrin does not only act as an attachment

receptor but it may also direct the virus to the clathrin-mediated

internalization route (Joki-Korpela et al., 2001).

Field isolates of FMDVs can recognize several integrins, including αvβ3

(Berinstein et al., 1995), αvβ6 (Jackson et al., 2000b), αvβ1 (Jackson et al.,

2002), and α5β1 (Jackson et al., 2000a). Binding of FMDVs to integrins is

RGD-specific (Jackson et al., 1997; Jackson et al., 2000b). However, if the

RGD sequence of FMDVs is mutated or if the viruses are grown in cell

cultures (Jackson et al., 1996; Sa-Carvalho et al., 1997), they are able to

switch the receptor to heparan sulphate on the cell surface (Baranowski et al.,

2000).

α2β1 integrin as a receptor for EV1The interaction of EV1 with α2β1 integrin (Bergelson et al., 1992) is serotype

specific as only EV1 and its homologue EV8 among echoviruses bind to the

integrin (Bergelson et al., 1993b; Ohman et al., 2001). In contrast to CAV9,

HPEV1 and FMDVs, EV1 does not carry an RGD tripeptide. α2β1 integrin

(VLA-2) is expressed in several cell types, including fibroblasts, platelets,

endothelial cells, and epithelial cells from multiple sites, i.e. skin,

gastrointestinal tract, lung and bladder (Zutter and Santoro, 1990). The natural

ligands of α2β1 integrin include collagen (Santoro, 1986), laminin (Elices and

Hemler, 1989) and E-cadherin (Whittard et al., 2002). After cell adhesion to

collagen, α2β1 integrin is known to regulate the mitogen-activated protein

kinase (MAPK) pathways (Heino, 2000). The α2 subunit contains a ligand-

binding I domain (α2I), which carries the C-helix not found in all αI domains

(Takada and Hemler, 1989; Bahou et al., 1994; Emsley et al., 1997) (Fig. 6).


27

The murine homologue of α2 integrin subunit does not bind to EV1 even

though it is 84% identical to human α 2 subunit (Edelman et al., 1994).

Instead, production of the human α2 subunit in rodent cells is required for

making these cells susceptible to EV1 infection, thus indicating that human

α2/mouse β1 heterodimers can serve as functional EV1 receptors (Bergelson

et al., 1993b; Zhang and Racaniello, 1997). However, when the murine

α2 subunit was replaced by human α2I domain, the murine α2β1 integrin also

supported virus binding (Bergelson et al., 1994b). The finding suggested that

EV1 binds to the α2I domain of α2β1 integrin, like a physiological ligand

collagen (Takada and Hemler, 1989; Bahou et al., 1994; Kamata et al., 1994).

This was further supported by a finding that a bacterial fusion protein of the

α2I domain specifically bound EV1 and prevented virus attachment to the

cells (King et al., 1995).

Figure 6. In the α 2I domain, one shortantiparallel and five parallel β-strands (βA-βE) form a core β -sheet, which issurrounded by seven amphipathic α-helices(Lee et al., 1995; Emsley et al., 1997). TheC-helix extends from a top of strand βE andcreates a groove, where collagen bindsthrough interactions with the MIDAS. Theresidues 199-201, 212-216 and Arg289 havebeen reported to interact with EV1 (King etal., 1997; Dickeson et al., 1999).

Even though both collagen and EV1 bind to the α2I domain, their interactions

with integrin differ in many aspects. In contrast to collagen, the virus binding

to α2β1 integrin does not discriminate between inactive and active

conformation of the integrin (Emsley et al., 2000). Studies with monoclonal

antibodies against α2I domain suggested that binding sites of EV1 and

collagen within the α2I domain are different (Kamata et al., 1994; Kamata

and Takada, 1994). In addition, the MIDAS site, essential for collagen


28

binding, is not involved in the interaction of EV1 and the integrin (King et al.,

1997). Thus, EV1-α2β1 integrin interaction is not dependent on any particular

divalent cation (Bergelson et al., 1993a).

The studies with murine and human α2I chimeras identified the EV1 binding

sites in the α2I domain as amino acids 199-201 and 212-216 (King et al.,

1997). These two regions interacted with the virus independently. All these

residues lie on the exposed face of the I domain (Fig. 6). To investigate

further the determinants of ligand binding specificity of α2β1, a collagen-

binding chimera of α2 and α1 I domains was constructed (Dickeson et al.,

1999). However, EV1 could not bind to α1I domain, and binding to α2I

domain was lost in a chimera containing the αC region, the αC-α6 loop, and

the α 6 helix of α 1. Further mutational analysis of the α1I/α2I domain

chimeras identified amino acid Arg289 in αC-α6 loop of α2I domain to be

critical for virus binding (Dickeson et al., 1999) (Fig. 6).

2.3 Other picornavirus receptors

Picornavirus receptors of Ig superfamily bind to the virus canyonSeveral picornaviruses, including PVs, major group HRVs and CBVs use

cell-surface molecules belonging to the immunoglobulin superfamily (IgSF)

as their receptors (Table 2). Picornavirus receptors of IgSF are type I

transmembrane glycoproteins and consist of tandem repeats of two to five Ig-

like domains, a transmembrane domain, and a short cytoplasmic tail (Fig. 4).

The interactions of picornaviruses with IgSF members are usually mediated

by the amino terminal domain (D1) of receptors (Rossmann et al., 2002).

PVR/CD155. Cell binding and entry of PVs 1-3 rely, as determined so far, on

one receptor, the poliovirus and vitronectin receptor PVR (CD155)

(Mendelsohn et al., 1989). PVR contains three extracellular Ig–like domains

and a short cytoplasmic tail (Koike et al., 1991). It shares significant

homology with nectins (Hogle, 2002), which are adhesion proteins related to

herpes virus entry (Geraghty et al., 1998). PVR interacts with the dynein light

chain that may direct its retrograde axonal transport in endocytic vesicles


29

(Mueller et al., 2002) and it is involved in NK-mediated killing of tumour

cells (Reymond et al., 2004).

Several lines of evidence suggest that the first N-terminal domain (D1) of

PVR is responsible for virus binding and infection (Koike et al., 1991). The

cytoplasmic domain of PVR is not involved in the PV entry process and thus

signalling may not be important for PVR behaving as a virus receptor (Koike

et al., 1991). Since the crystal structure of PVR has not yet been solved, the

differences in the interpretation of the homology-built models of PV-receptor

interactions exist. Several authors have suggested that the D1 of PVR binds to

both outer (=south) and inner (=north) walls of the viral canyon and interacts

also with the floor of the canyon (Belnap et al., 2000b; He et al., 2000). Thus,

the receptor seems to bridge the virus canyon. In contrast, Xing and co-

authors claimed that PVR does not interact with the bottom of the canyon.

They have described the PVR as boot-like in shape, the tip of the foot

contacting the south wall of one protomer and the heel laying on a protrusion

in the neighbouring one (Xing et al., 2000).

ICAM-1. The major group HRVs, such as HRV14 and HRV16 (Greve et al.,

1989; Staunton et al., 1989; Tomassini et al., 1989), and CAV21 (Shafren et

al., 1997) interact with an intercellular adhesion molecule-1 (ICAM-1).

ICAM-1 contains five Ig-like domains. It regulates leukocyte adhesion, and

its natural ligands are integrins (Staunton et al., 1988). In cryo-electron

microscopy (cryo-EM) and X-ray crystallography studies, the D1 domain of

ICAM-1 was found to interact primarily with the floor and south wall of the

HRV canyon (Olson et al., 1993; Bella et al., 1998; Kolatkar et al., 1999).

The orientation of the ICAM-1 molecule with respect to the virus surface is

almost the same between HRV14 and HRV16 but different from CAV21

(Xiao et al., 2001; Rossmann et al., 2002).

CAR. A coxsackie-adenovirus receptor (CAR) functions as a receptor for the

six CBV serotypes as well as for certain adenoviruses (Bergelson et al., 1997;

Tomko et al., 1997). CAR is a broadly distributed type I membrane

glycoprotein of the Ig-family and it has only two Ig-like extracellular

domains, thus being shorter than PVR or ICAM-1 (Tomko et al., 1997). It is a


30

component of the tight junctions and thus it is involved in regulating the

passage of macromolecules and ions across cell monolayers (Cohen et al.,

2001). CBVs and adenoviruses bind to different but overlapping sites of

domain 1 in CAR (He et al., 2001). Human, mouse and zebrafish CAR can

bind to CBV3, and conserved residues in the CBV3 footprint suggest that the

binding is clustered in the canyon region (Bergelson et al., 1998; Petrella et

al., 2002). Also, cryo-EM reconstruction of the full-length CAR in complex

with CBV3 showed that the D1 domain of the receptor binds to the CBV3

canyon (He et al., 2001). The cell-associated and soluble forms of CAR can

induce the conformational alterations in the viral capsid (Milstone et al.,

2005).

The binding of the IgSF-like receptor to a canyon of picornaviruses can lead

to conformational changes in the viral capsid, as studied most extensively

with PVs (Hogle, 2002). PV can undergo structural alterations when it is

exposed to cell-associated or soluble receptors (Arita et al., 1998). These

changes in virus conformation have been studied using cryo-EM

reconstruction, X-ray crystallography and CsCl- or sucrose gradient

centrifugation methods. In linear sucrose gradient ultracentrifugation, the

different conformations of viral capsids are sedimented based on their

velocity. The sedimentation is expressed as Svedberg’s coefficient of 160S for

intact capsid, 135S for capsid lacking VP4 and 80S for capsid lacking VP4

and the genomic RNA (Hogle, 2002). However, the sedimentation

coefficients may vary between different picornaviruses, and, moreover, some

forms of viral capsids might be too unstable to be recognized by this method.

As proposed in a current model for PV uncoating and host cell entry, the

receptor acts as a catalyst, which, after PV binding, is able to trigger the

externalisation of VP4 and the N termini of VP1. As a result, N-terminal

helices of VP1 are inserted into and rearranged in the lipid membrane,

resulting in a pore. The viral RNA can be released into the cytoplasm through

the pore when a plug formed by VP3 is removed (Belnap et al., 2000a).

Whether this pore is formed in the plasma membrane or in the membrane of

intracellular membranous vesicles is still unknown.


31

Other examples of picornavirus receptorsDAF. A complement regulatory protein, decay accelerating factor (DAF;

CD55) acts as a receptor for several echovirus serotypes (Bergelson et al.,

1994a; Ward et al., 1994; Clarkson et al., 1995; Powell et al., 1998) and

enterovirus 70 (Karnauchow et al., 1996) as well as a secondary receptor for

CAV21 (Shafren et al., 1997) and CBVs 1, 3, and 5 (Bergelson et al., 1995;

Shafren et al., 1995)(Fig. 4, Table 2). DAF is expressed in most mammalian

cells and it consists of four short consensus repeats (SCRs) and a

glycosylphosphatidylinositol anchor (GPI) (Medof et al., 1987). EV7

(Clarkson et al., 1995; He et al., 2002) and CBV3 (Bergelson et al., 1995)

bind to a region near or in the third SRC domain, but enterovirus 70

(Karnauchow et al., 1998) and CAV21 (Shafren et al., 1997) interact with the

first SRC domain. In EV7, DAF binds around the two-fold axes (He et al.,

2002) and it cannot induce conformational changes in the viral capsid (Powell

et al., 1997). Accordingly, interaction with DAF is not sufficient to initiate the

conformational alterations of CBV3 (Milstone et al., 2005). Therefore, DAF

interactions with enteroviruses differ remarkably from IgSF receptor-

picornavirus interactions.

VLDL-R. The minor group HRVs bind to a very-low-density lipoprotein

receptor (VLDL-R) (Hofer et al., 1994), which normally shuttles the VLDL-

particles into the host cell. The binding site of VLDL-R in HRV2 structure is

on a small, star-shaped dome around the five-fold axes (Hewat et al., 2000).

The binding of HRV2 to its receptor leads to clathrin-mediated endocytosis,

followed by uncoating under conditions of low endosomal pH (Bayer et al.,

2001).

Heparan sulphate. The field isolates of FMDVs, which normally bind to

integrins (Berinstein et al., 1995), can adapt to use heparan sulphate as a

receptor in laboratory cell cultures (Jackson et al., 1996; Neff et al., 1998).

The binding site of heparan sulphate is a shallow depression on the virus

surface, located at the junction of VP1, VP2 and VP3 (Fry et al., 1999).

Heparan sulphate may also be involved in attachment of certain echoviruses

on the cell surface (Goodfellow et al., 2001).


32

Sialic acid(s) may act in enterovirus 70 binding and productive infection

(Alexander and Dimock, 2002). This has been confirmed by Haddad et al.,

who suggested that enterovirus 70 is able to use sialyted receptors other than

DAF in cultured human leukocyte cell lines (Haddad et al, 2004). Also, the

presence of sialic acid on cellular receptors is required for HRV87 attachment

and infection (Uncapher et al., 1991).

ββββ2 microglobulin. On the cell surface, β2m is expressed in association with

MHC class I molecules. Antibodies against β2m have been found to prevent

many enterovirus infections, suggesting that β2m could act as a secondary

receptor for many serotypes of echoviruses, including EV1 (Ward et al.,

1998) as well as for some coxsackieviruses (Triantafilou et al., 1999).

However, the exact role of β2m in virus binding has remained unclear and it

may, more probably, have an indirect role in virus-cell interactions.

3. Internalization of picornaviruses into host cells

The aim for the virus is to enter a suitable site, either the cytoplasm or the

nucleus of the host cell, for replication. Most nonenveloped viruses use

endocytosis which carries them into the host cell through the membrane

barrier and cortical actin network (Fig. 7). For a long time, clathrin-mediated

endocytosis was regarded as the main entrance route. More recently,

investigations have led to recognition of other significant endocytic pathways,

including lipid rafts, caveolae, macropinocytosis and non-caveolae and non-

clathrin -dependent mechanisms (Johannes and Lamaze, 2002). It can be

expected that additional endocytic pathways will be identified in the future

(Damm et al., 2005; Kirkham et al., 2005). The data accumulated on virus

endocytosis indicate that many viruses can switch from one uptake

mechanism to another and simultaneously utilize several endocytic routes in

order to enter efficiently into the host cell (Sieczkarski and Whittaker, 2005).

So far, picornaviruses have been reported to use clathrin-, caveolae-, and lipid

raft- dependent uptake mechanisms. Instead, PV may protrude its genome into


33

the host cell directly from the plasma membrane by generating a pore into the

membrane (Hogle, 2002), as explained in the Chapter 2.3.

Figure 7. Endocytic mechanisms of virus entry. In addition to endocytic uptakemechanisms, nonenveloped viruses, such as PV, may release their genome into thecytoplasm through a pore, formed at the plasma membrane. Some enveloped virusesintroduce their genome into the host cell via direct membrane fusion events.

Viruses have served as excellent tools for the studies of the cellular endocytic

mechanisms since their uptake is easier to follow compared to smaller natural

ligands (Pelkmans and Helenius, 2003). On the other hand, the lack of

specific markers of different entry pathways has made it challenging to define

the uptake routes for viruses. Inhibitors of cellular functions are efficient but

rather inaccurate in studies of endocytosis. Usually, dominant-negative

mutants derived from both structural and regulatory proteins of endocytic

machinery are more specific than chemical inhibitors (Sieczkarski and

Whittaker, 2002; Pelkmans and Helenius, 2003). However, in some cases the

results obtained with dominant negative mutant proteins have not been

interpreted correctly. For example, dominant negative dynamin 2 was thought


34

to affect only the clathrin route but later studies have revealed its inhibitory

effect on other routes of entry, including caveolar endocytosis (Damke et al.,

1994; Henley et al., 1998; Oh et al., 1998).

Microscopical methods are certainly required for detailed studies on endocytic

routes. The (immuno)electron microscopy (EM) has been widely used to

define the morphology of intracellular structures involved in virus uptake

(Helenius et al., 1980; Kartenbeck et al., 1989). Prelabelled cellular

molecules, characterized to utilize a certain endocytic pathway, as well as

specific antibodies against cellular proteins have been applied in

immunofluorescence microscopy. Most recently, real-time microscopy of

living cells has provided a sophisticated method to follow the trafficking of

fluorescently tagged or prelabelled proteins and viruses, such as adenovirus

(Suomalainen et al., 1999), simian virus 40 (Pelkmans et al., 2001) and

influenza virus (Lakadamyali et al., 2003). The approaches reviewed briefly

above have been applied in this thesis to the studies of endocytosis of EV1.

3.1 Clathrin -mediated endocytosis

Clathrin-coated pits are formed from a basketlike framework of clathrin

(Kirchhausen, 2000). Several proteins, including adaptor complex AP-2 and

dynamin GTPase, regulate the assembly and fission of the pits (Takei and

Haucke, 2001). Upon ligand binding to its receptor in the clathrin-coated pits,

the pits pinch off to form intracellular clathrin-coated vesicles (Brodsky et al.,

2001). Within seconds, clathrin-coated vesicles shed their coat and fuse with

early endosomes (EEs). After 5 to 15 minutes, EEs fuse with late endosomes

(LE) that have a more acidic pH. Endocytosed receptors do not always reach

the LEs but, instead, they can be recycled back to the cell surface in

approximately 20 min. Some molecules are transported from LEs into

lysosomes, which contain hydrolytic enzymes for degradation.

Many viruses, including adenoviruses, alphaviruses, hantaviruses,

orthomyxoviruses, parvoviruses, and some picornaviruses (DeTulleo and

Kirchhausen, 1998; Marsh and Pelchen-Matthews, 2000) take advantage of

clathrin-mediated endocytosis.


35

The clathrin-mediated endocytosis of the minor group HRVs, especially

HRV2, is probably the most extensively studied entry mechanism among

picornaviruses (Prchla et al., 1994). The recent report on HRV2 entry showed

that the infection can be inhibited by the dominant negative mutant of

dynamin 2 GTPase as well as by more specific dominant-negative inhibitors

of clathrin-mediated endocytosis, such as the SH3 domain of amphiphysin or

the C-terminal domain of AP180 (Snyers et al., 2003). However, some studies

have challenged the endocytosis of HRV2 via clathrin-coated pits (Bayer et

al., 2001; Huber et al., 2001).

The endocytosis of HRV2 is mediated by the VLDL-R, which is dissociated

from the virus in EEs (Brabec et al., 2003). The virus reaches LEs, where it

undergoes conformational alterations, dependent on low pH (Bayer et al.,

2001; Huber et al., 2001). Upon uncoating, the viral RNA is transferred into

the cytoplasm across a pore in the endosomal membrane and viral capsid

proteins may be transported to lysosomes for degradation (Prchla et al., 1994;

Prchla et al., 1995; Schober et al., 1998).

The entry of major group HRVs, such as HRV14, proceeds also via clathrin-

coated pits to EEs (Schober et al., 1998). Based on some studies, the entry

and uncoating of HRV14 appears to be pH-dependent (Grunert et al., 1997;

Nurani et al., 2003). However, it has also been suggested that the virus could

cause a lytic disruption of EEs by a receptor-dependent manner in the absence

of endosomal acidification (Bayer et al., 1999). This could lead to release of

viral RNA into the cytoplasm (Schober et al., 1998; Bayer et al., 1999).

HPEV1, which binds to αv integrins on the cell membrane, enters the cells

through clathrin-mediated endocytosis (Joki-Korpela et al., 2001). The

receptor, αvβ3 integrin, does not colocalize intracellularly with the virus.

After 5 min of internalization, the virus is in EEs, and after 30 min, it is found

in the LEs. After 30-60 min, the capsid proteins are located in both the

endoplasmic reticulum (ER) and the cis-Golgi network. The viral RNA may

be released early during the entry process, since depolymerization of


36

microtubules did not block viral infection even though it inhibited movement

of HPEV1 capsid proteins to LEs (Joki-Korpela et al., 2001).

A recent study of internalization and trafficking mechanisms of CAR-

dependent strain of CBV3 revealed that the virus enters cells via clathrin-

coated pits (Chung et al., 2005). The virus was detected in the pits and in

clathrin-coated vesicles by immuno-EM. CBV3 was also found in clathrin-

coated pits and vesicles and in EEs in immunofluorescent labelling of infected

cells. Moreover, endosomal acidification and dynamin 2 were shown to be

essential factors for the infection (Chung et al., 2005).

In addition to direct uncoating on the cell membrane, clathrin-mediated

endocytosis has been suggested for PVs (Zeichhardt et al., 1985;

Willingmann et al., 1989; Kronenberger et al., 1998). However, PV infection

is not obligatorily dependent on dynamin 2 (DeTulleo and Kirchhausen,

1998), a marker of clathrin- and caveolae-mediated pathways. More recently,

PV infection was shown to be inhibited in the presence of cholesterol

depletion. However, the virus did not localize in the detergent-insoluble

microdomains (Danthi and Chow, 2004). Therefore, it was suggested that PV

infection and the release of viral RNA are dependent on cholesterol but not on

lipid rafts. Thus, further investigations are required to clarify the entry

mechanisms of PVs.

3.2 Lipid raft -mediated endocytosis

Lipid rafts are membrane microdomains, enriched in cholesterol, specific

glycosphingolipids and several signalling molecules (Simons and Ikonen,

1997), as well as integrins (Brown, 2002). Rafts are rather small dynamic

structures that are stabilized through interactions with the cytoskeleton

(Kenworthy, 2002). In response to various stimuli, lipid rafts can aggregate

into larger platforms. Cell surface caveolae are highly specialized type of lipid

rafts, and they contain caveolin-1 as their main protein component

(Kurzchalia and Parton, 1999). Even though several studies during last


37

decades have focused on the lipid rafts, their size, dynamics and composition

are still under debate (Mayor and Rao, 2004).

Endocytosis via lipid rafts can be dependent on or independent of dynamin 2,

but otherwise, this internalization pathway is rather poorly known (Pelkmans

and Helenius, 2003). However, recent studies on alternative entry pathways of

simian virus 40 (SV40) and cholera toxin (CTX) may illuminate the raft-

dependent entry mechanisms (Damm et al., 2005; Kirkham et al., 2005).

Rafts are not only involved in viral endocytosis but also in the assembly and

budding of viruses (Suomalainen, 2002). Viruses that may use rafts or raft-

located receptors for the endocytosis and/or fusion events include group A

rotaviruses (Isa et al., 2004), avian sarcoma and leucosis virus (Narayan et al.,

2003), certain enteroviruses (Stuart et al., 2002a; Triantafilou and

Triantafilou, 2003, 2004), SV40 (Damm et al., 2005), and HIV (Manes et al.,

2000).

Recently, EV11 was reported to be endocytosed via lipid rafts and/or

caveolae (Stuart et al., 2002a). Interestingly, the EV11 mutant that does not

bind to DAF, a receptor for EV11, was not internalized through rafts,

indicating that internalization is specifically directed by the receptor. Indeed,

DAF contains a GPI-anchor, and it is found in plasma membrane lipid rafts

(Bergelson et al., 1994a; Mayor et al., 1994). EV11 is also able to infect cells

not expressing caveolin-1. In such a cell line, similarly to the cell line

expressing caveolin-1, the virus was copurified with detergent insoluble

membrane microdomains within 30 min-1 h p.i. Thus, EV11 may use both

caveolae and non-caveolar lipid rafts in its entry process. Also, an intact actin

cytoskeleton and microtubule network are required prior to uncoating of the

virus (Stuart et al., 2002a).

A recent paper suggested that lipid raft microdomains could also play a role in

the entry process of CAV9 (Triantafilou and Triantafilou, 2003). The viral

receptor, αvβ3 integrin as well as possible accessory molecules, such as

GRP78 and MHC class I molecule, were found in increased concentrations in

lipid rafts when studied at 30-60 min after CAV9 binding to cells. Moreover,

raft-interfering agents inhibited CAV9 infection. Also, another enterovirus,


38

CBV4, was reported to use rafts for efficient entry into the Golgi (Triantafilou

and Triantafilou, 2004). However, the role of caveolae cannot be ruled out in

the endocytic processes of these viruses because it was not examined.

3.3 Caveolae -mediated endocytosis

The name of caveolae is derived from Latin and means little caves (Palade,

1953). Caveolae are flask-shaped invaginations in cell membranes, formed

from a structural coat protein caveolin-1 (Rothberg et al., 1992; Schlegel and

Lisanti, 2001). Cholesterol binds to caveolin and seems to be necessary for

the structure of caveolae (Murata et al., 1995; Ikonen and Parton, 2000).

Moreover, cholesterol is involved in signalling and trafficking functions of

caveolae. Caveolae contain several signalling proteins, such as protein kinase

C (Mineo et al., 1998), GTP binding proteins and non-receptor tyrosine

kinases (Sargiacomo et al., 1993; Lisanti et al., 1994). Caveolae are proposed

to act as "organized transducing centres that concentrate key signalling

molecules in a compartment to create rapid, efficient and specific

transmission of signals from the cell surface into the cell" (Lai, 2003). On the

other hand, caveolae contain molecular machinery for regulated, receptor-

mediated endocytosis and transcytosis of selected ligands via vesicle budding,

docking and fusion (Schnitzer et al., 1995).

Caveolae are used in the entry of natural ligands (e.g. albumin)(Schnitzer et

al., 1994), toxins (e.g. cholera toxin; CTx)(Montesano et al., 1982; Nichols,

2002), bacteria (e.g. E.coli) (Shin et al., 2000) and viruses (Pelkmans and

Helenius, 2002). The binding of a ligand to a receptor in caveolae may trigger

the internalization of caveolae. The GTPase dynamin, depolymerization of

actin and specific signalling events are required for the internalization (Parton

et al., 1994; Henley et al., 1998). The internalized caveolar vesicles of 60-70

nm can deliver the cargo to cytoplasmic organelles, caveosomes (Pelkmans et

al., 2001). In addition, when recruited by small GTPases such as Rab5, the

caveolar vesicles can fuse with other organelles such as EEs and form distinct

and stable membrane domains (Pelkmans et al., 2004).

Caveosomes have been defined as pre-existing organelles, enriched in

caveolin-1 and devoid of markers of classical endocytic pathways and


39

biosynthetic organelles, including Rab GTPases 4, 5, 6, 9, and 11, and

endosomal and lysosomal markers (Pelkmans et al., 2001; Pelkmans and

Helenius, 2002, 2003). A significant difference with endosomes is the pH-

neutrality of caveosomes (Pelkmans et al., 2001). The ligands may be

transported to caveosomes also directly from lipid rafts by a mechanism that

does not involve cell surface caveolae, caveolar vesicles or clathrin-coated

vesicles (Damm et al., 2005; Kirkham et al., 2005). Caveolar vesicles and

caveosomes can further deliver their cargo to the endosomes, Golgi,

endoplasmic reticulum (ER) or lysosomes, often through microtubule-directed

movements (Conrad et al., 1995; Nichols, 2002).

Viruses using caveolar endocytosis. Simian virus 40 (SV40) was the first

virus characterized to use caveolae-mediated endocytosis (Anderson et al.,

1996). Many other viruses, e.g. EV1 (Marjomäki et al., 2002), filoviruses

(Empig and Goldsmith, 2002), some coronaviruses (Nomura et al., 2004) and,

in some cases, polyomaviruses (Richterova et al., 2001; Eash et al., 2004;

Gilbert and Benjamin, 2004) can also enter the cells via caveolae. So far, EV1

is the only picornavirus shown to utilize caveolar endocytosis, as discussed in

the Results and Discussion of this thesis. Because the uptake of SV40 to

caveosomes has been investigated extensively, it is reviewed here in detail

(Pelkmans and Helenius, 2003) (see Fig. 10., p.67)

SV40 is a nonenveloped DNA virus and it has an icosahedral capsid of 50 nm.

The capsid is composed of 360 subunits of the major coat protein VP1 and

containing minor amounts of VP2 and VP3. On the cell surface, SV40 binds

to MHC class I molecule (Anderson et al., 1996; Stang et al., 1997) and

ganglioside GM1, located into lipid rafts (Tsai et al., 2003). Within 20

minutes, the mobile virus particles are trapped into caveolae (Pelkmans et al.,

2001). As a consequence of SV40 binding to its receptor, intracellular

caveolar vesicles are recruited to the membrane for formation of additional

caveolae. The MHC class I molecule is transported from non-caveolar region

to the cell surface caveolae upon the virus binding but the receptor is not

endocytosed with a virus (Stang et al., 1997; Anderson et al., 1998).


40

However, the ganglioside GM1 may be transported with SV40 from the

plasma membrane to the ER (Tsai et al., 2003).

In caveolae, SV40 triggers a signal transduction cascade (Chen and Norkin,

1999) that leads to actin cortex depolymerization, local tyrosine

phosphorylation and production of phosphatidyl 4,5-bisphosphate (Pelkmans

et al., 2002; Pelkmans and Helenius, 2003). Subsequently, both actin and

dynamin are recruited to caveolae and actin tails are formed at the site of

caveolae internalization (Pelkmans et al., 2002). These events lead to

relatively slow but efficient internalization of virus-containing caveolae to the

cytosol.

A recent paper reported that some of incoming SV40 colocalizes with Rab5a,

a GTPase normally located into EEs (Pelkmans et al., 2004). When a

dominant active-mutant of Rab5a was expressed in the cells, SV40 was

trapped into a greater extent into endosomes, which also contained caveolin-1.

Even though caveolar vesicles, which carry SV40 particles, transiently

interact with EEs also under normal conditions, this pathway is not obligatory

for SV40 infection (Pelkmans et al., 2004).

Virus-carrying caveolar vesicles fuse with caveosomes after the next 40 min

to 3 h, making caveosomes increasingly dynamic (Pelkmans et al., 2001). A

recent paper reported that SV40 can, in mouse knock-out cells lacking

caveolin-1, enter the caveosome-like organelles via an alternative pathway

that is not dependent on caveolin-1 or dynamin 2 but involves tyrosine

phosphorylation events (Damm et al., 2005). In contrast to caveolar uptake,

this pathway appears to be more rapid and results in removal of the virus from

cell surface lipid rafts into caveosome-like organelles through non-caveolar,

non-clathrin -coated vesicles. Interestingly, this entry pathway may also be

utilized by SV40 in cell lines that contain caveolin-1 (Damm et al., 2005).

From caveosomes, SV40 is carried to the ER in long, tubular vesicles that

move along microtubules (Pelkmans et al., 2001). The majority of virus

particles remain in the smooth ER for up to 16 h or longer (Norkin et al.,

2002). The details concerning the transport of viral particles to the nucleus for

replication remain unclear.


41

4. Host cell gene expression during picornavirus infection

Viruses are able to cause dramatic changes in host cells in order to maximise

the efficiency of their replication and to overcome the host cell defence

mechanisms. The modifications of cellular functions can be brought about by

interactions of virus with its receptor, by viral replication and by direct

interference with the functions of cellular proteins and organelles (Knipe et

al., 2001).

To obtain optimal conditions for replication, picornaviruses can take over the

host cell translation machinery and cause inhibition of host cell protein

synthesis (shut-off). Moreover, cellular transcription can be inhibited and host

cell gene expression pattern altered during the infection (Racaniello, 2001).

At the molecular level, these changes result in the activation of a variety of

cellular pathways, including those related to stress response, cytokine

production and apoptosis (programmed cell death). In addition, picornaviruses

can remodify cellular membranes, interfere with intracellular protein

trafficking and alter numerous other cellular functions (Carrasco et al., 2002).

At the end of the picornavirus replication cycle, the exploitation of host cell

machinery is usually observed as a cytopathic effect (CPE), which includes

cell rounding and detachment and, finally, cell death (Racaniello, 2001).

Picornaviruses can also exhibit apoptotic activity to release the progeny viral

particles from the host cell and, on the other hand, they can induce

antiapoptotic activities to prevent premature cellular death (Tolskaya et al.,

1995). All these events in host cells during the infection affect the viral

pathogenesis in the target tissue.

4.1 cDNA array studies of host cell gene expression in enterovirusinfection

The traditional approaches, such as Northern blot analysis, have been

successfully used in measuring changes in cellular mRNA expression during

viral infections. More recently, cDNA (micro)array techniques, which allow

the simultaneous determination of the levels of thousands of specific mRNA


42

species, have been applied in the characterization of host cell gene expression

during virus infections (Jenner and Young, 2005).

cDNA arraysFor cDNA array analysis, RNA samples can be extracted from tissues or from

cultured cells. The samples are reverse-transcribed into cDNA in the presence

fluorescently or radioactively labelled nucleotides. cDNAs are then hybridised

either onto membranes or microarrays, carrying the cDNA clones (cDNA

membrane arrays, cDNA microarrays) or oligonucleotide sequences (Duggan

et al., 1999). The signal is detected either by measuring the fluorescence

intensities or radioactivity of the hybridised spots (Fig. 8).

Figure 8. The general principles of the cDNA array technique, used for screening ofthe host cell gene expression during virus infection. The total RNA samples areisolated from uninfected control cells and infected cells for synthesis of radioactivelylabelled cDNAs. cDNA transcripts are then hybridised to parallel membranes, whichhave been spotted with hundreds or thousands of cDNA clones.

The ability to screen a wide variety of host cell genes with cDNA arrays

provides an efficient method to clarify gene alterations during viral

pathogenesis (Jenner and Young, 2005). In addition, the method is relatively

fast when considering the amount of data that is generated (Brown and


43

Botstein, 1999). However, there are also some drawbacks in the technique.

Reproduction of the results is difficult because the phase of cell cycle, quality

of input virus, and temperature changes during collection of samples may

vary between the parallel experiments. Sensitivity of the cDNA (micro)array

system is dependent on many factors, such as the quality of samples, and it

may also vary between the experiments, leading to wrong interpretation of the

results. The selection of the most reliable control genes (house-keeping genes)

is challenging in studies of virus infections, because the infection may result

in altered expression of normally "stable" genes. Also, the tools for "data

mining", such as statistical analysis and interpretation of the data, are critical

(Bassett et al., 1999), even though profiling of affected genes into functional

clusters has greatly facilitated the elucidation of results.

Many other factors than those observed at the gene expression level may

contribute to the onset and severity of the disease in pathogenetic models, and

thus, not too far-reaching conclusions from the results of cDNA array analysis

should be drawn. Despite these concerns, cDNA array screening may help to

explain how the host cell "sees" a virus at gene expression level and, on the

other hand, how viruses exploit the host cell machinery (Manger and Relman,

2000).

Studies of enterovirus infection with cDNA arraysThe first cDNA array study of picornavirus infection concentrated on the

identification of eucaryotic mRNAs that are translated in the presence of PV-

induced shut-off of host cell protein synthesis (Johannes et al., 1999). Here,

the cellular mRNAs associated with polysomes of PV-infected HeLa cells

were hybridised to the cDNA microarrays (Johannes et al., 1999). Among the

7000 genes studied, around 0.3% of genes showed changes over 1.7-fold at 3

h p.i. The genes enriched in polysomes were encoding, for example, for

"immediate early" genes, proteins of mitogen-activated-protein kinase

(MAPK) pathways, oncogenes, DNA binding proteins and cellular receptors.

Other cDNA array studies of enterovirus infections have focused on CBV4 -

induced pancreatitis in mice (Ostrowski et al., 2004), and on the pathogenesis

of CBV3 -induced myocarditis in mice (Taylor et al., 2000). Of the 7000

genes screened, approximately 2% had changed levels of expression during


44

the CBV3 infection in the myocardium (Taylor et al., 2000). The affected

genes were clustered into functional groups of host defence, cell signalling,

cell division, cell structure/motility, protein and gene expression, cell

metabolism and mitochondrial genes. In addition to pathogenic mouse

models, cDNA arrays have been applied to the studies of CBV3 infection in

HeLa cells (McManus et al., 2002). Even though the host cell gene expression

patterns are somewhat different during enterovirus infections, common

clusters of activated genes can be found, as discussed below.

Immediate early (IE) genes. The expression of IE genes is rapidly and

transiently induced as a response of extracellular stimuli to alter patterns of

cellular gene expression (Sng et al., 2004). The expression of these genes is

dependent only on modification of factors already present in cells (Thomson

et al., 1999). IE genes encode for chemo-attractants, cytoplasmic enzymes,

ligand-dependent transcription factors and inducible transcription factors,

such as fos (c-Fos, FosB, Fra-1, Fra-2), jun (c-Jun, JunB, JunD) and

activating transcription factors (ATF) (e.g. ATF2, ATF3). The gene products

of these families are the main components of transcription factor AP-1 that

controls the transcription of genes involved in cell proliferation,

transformation, survival and death (Shaulian and Karin, 2002).

In the cDNA array analysis of PV-infected cells, several IE genes were found

to be enriched in polysomes and their expression was increased compared to

the uninfected control cells (Johannes et al., 1999). The upregulated IE genes

included Pim-1 proto-oncogene, c-myc proto-oncogene, ATF3, and

transforming growth factor-β-inducible early growth response protein

(TIEG). Correspondingly, ATF3 as well as IE genes c-jun and c-fos have been

found to be transiently overexpressed at early time points in CBV3-infected

HeLa cells (McManus et al., 2002). In addition, gene profiling of CBV3-

infected mouse hearts has revealed an increased expression of a related gene,

ATF4, during the inflammatory phase of infection (Taylor et al., 2000). ATF3

and ATF4 are transcriptionally upregulated in most of the cell stress responses

(Rutkowski and Kaufman, 2003). They can either act as transcriptional

repressors or bind to c-jun in order to facilitate cellular transcription.


45

Genes encoding proteins of MAPK pathways. The individual IE genes as

well as AP-1 are regulated by mitogen-activated protein kinases (MAPKs),

such as extracellular signal-regulated kinase (ERK) -1/2, the Jun-N-terminal

kinase/stress-activated protein kinase (JNK/SAPK), p38 MAPK and ERK5

(Chang and Karin, 2001). ERK pathways mainly respond to mitogens and

growth factors, whereas JNK/SAPK and p38 are activated by environmental

changes and inflammatory cytokines. MAPKs mediate apoptosis, cell

transformation and responses to cell stress and cytokines (Chang and Karin,

2001).

The activation of MAPK pathways at the protein level has been observed

during several enterovirus infections, including EV11, EV12, CBV3 (Huber,

1999) and EV1 (Huttunen et al., 1998) infections. The activation of ERK1/2

during CBV3 infection leads to virus-induced CPE and apoptosis (Huber,

1999; Luo et al., 2002). The regulators of MAPK pathway, such as

MAPKK3b gene, were associated in increased amounts in polysomes in PV-

infected cells in the cDNA microarray study (Johannes et al., 1999). The

authors speculated that the activation of p38 MAPK pathway by MAPKK3b

during PV infection could, for example, lead to production of inflammatory

cytokines.

IRES -containing genes. In the presence of a complete enterovirus-induced

inhibition of host cell protein synthesis (=shut-off), only IRES-containing

viral RNA or host cell mRNAs that carry an IRES can be translated (Pelletier

and Sonenberg, 1988). The shut-off effect is mainly caused by a cleavage of

the translation initiation factor eIF4GI/II by enteroviral protease 2A (Knipe et

al., 2001). After the cleavage, eIF4G cannot attach via eIF4E to the 5' capped

structures in the cellular RNAs to initiate translation. Pim-1 and c-myc mRNA

levels increased during PV infection in HeLa cells. Both of these genes

contain an IRES element, which enables their translation in the presence of

PV-induced shut-off. Pim-1 encodes a serine-threonine protein kinase that can

cooperate with c-myc during cellular transformation (Wang et al., 2001). c-

myc IRES has been shown to be activated during the mitosis and via MAPK

p38 during apoptosis (Stoneley et al., 2000).


46

4.2 The effects of EV1 on host cell gene expression

Transcription of IE genes c-jun, junB, and c-fos has been demonstrated to

increase after 5-10 h of EV1 infection by Northern blot analysis of the

infected cells (Huttunen et al., 1997) and by analysis of the transcription rates

of the genes (Huttunen et al., 1998). Activation of both p38 MAPK and ERK

1/2 pathways led to increased c-fos expression whereas p38 MAPK was the

main inducer of junB expression (Huttunen et al., 1998).

In these studies, it was of interest whether the virus-integrin interaction could

lead to IE gene activation, which is observed as a consequence of type I

collagen binding to the α2β1 integrin (Rana et al., 1994). As a result, EV1-

receptor interactions did not seem to influence the regulation of IE genes

during the infection. However, post-attachment events were required for IE

gene activation (Huttunen et al., 1997). This idea was supported by the fact

that the IE genes were not induced in EV1-infected cells in the presence of

antiviral compound WIN, which interferes with receptor binding and

uncoating of the virus. In addition, the transfection of viral RNA into the cells

was sufficient to induce the activation of junB (Huttunen et al., 1997).

Moreover, the induction of IE genes was described not to be unique for EV1-

α2β1 integrin interaction because a similar activation pattern of IE genes was

observed in cells infected with EV7 and with PV1, which use another receptor

for cell surface binding (Huttunen et al., 1997).

However, according to a recent study by Upla and colleagues, the interaction

of EV1 with α2β1 integrin is able to trigger signalling events, such as PKCαphosphorylation and activation of MAPK ERK1/2 (Upla et al., 2004). This is

in agreement with the fact that binding of natural ligands to integrins can lead

to regulation of the MAPK pathway(s) (Juliano et al., 2004). Interestingly, the

activation of ERK1/2 during CBV3 infection (Huber, 1999; Luo et al., 2002)

may also lead to the expression of IE genes. The early activation of ERK1/2 is

triggered by cell-surface binding and internalization of CBV3, but the late

activation requires virus replication. The similar two-step activation of

MAPKs could also occur in EV1 infection, thus explaining the somewhat

contradictory results (Huttunen et al., 1997, 1998; Upla et al., 2004).

Aims of the study

47

AIMS OF THE STUDY

The initial aims of this thesis were to investigate the interactions of EV1 with

α2β1 integrin, to characterize the endocytic uptake mechanism(s) for EV1,

and to study the host cell gene expression during enterovirus infection.

SPECIFIC AIMS:

Interactions of EV1 with αααα2ββββ1 integrin:• To study the attachment of EV1 to the cell surface and to α2β1

integrin• To study conformational changes and uncoating of EV1 as a

consequence of the cell surface/receptor binding• To produce the intact EV1 for structural cryo-EM studies of the

EV1-α2I domain complex• To define the binding sites of α2I domain in EV1 capsid

Endocytic uptake mechanisms for EV1:• To define how EV1 enters the host cell• To characterize the role of caveolae and caveosomes in EV1

internalization process• To illustrate the endocytic route of EV1 into caveosomes in more

detail using fluorescently labelled virus in real-time livemicroscopy

Intracellular effects of EV1 infection:• To screen the alterations in cellular gene expression during EV1,

CBV4 and PV1 infections by using a new cDNA array method

Materials and Methods

48

MATERIALS AND METHODS

The original publications in which the methods have been employed are indicated in

brackets. The immuno-electron microscopy (EM) (I), solid phase binding assays (I)

and cryo-EM-modelling of EV1-α2I domain structure (I) were completely carried out

by the collaborators, and are explained in detail in original publications I and II.

1. Viruses (I-IV)Viruses. EV1 (Farouk strain), CBV4 (JVB), PV1 (Sabin) and SV40 were originally

obtained from the American Type Culture Collection (ATCC). EV1 was purified as

originally described by Abraham and Colonno (Abraham and Colonno, 1984).

Briefly, the infected cells and the supernatant were collected, and after three freeze-

thaw cycles, the virus was precipitated by PEG/NaCl and purified by

ultracentrifugation in 5-20% sucrose gradients. Purified CBV4 and PV were obtained

from M. Roivainen (National Public Health Institute, Finland). Purified SV40 was

obtained from L. Pelkmans (Swiss Federal Institute of Technology Zürich,

Switzerland)(Pelkmans et al., 2001). MOIs of 1-20 of purified viruses were used in

most of the experiments.

Radioactive labelling of viruses. To obtain radioactively labelled EV1 and PV1, the

infected cells were incubated in the presence of [35S]-methionine (50 µCi/ml;

Amersham Pharmacia Biotech) in Eagle minimal essential medium (MEM) deficient

in L-methionine (MEM-met) (GibcoBRL, Life Technologies) and purified as above.

Fluorescent labelling of EV1. The fluorescent labelling of purified EV1 (100-200 µg

of virus), with a ten times higher molar concentration of Texas Red-X succinimidyl

ester (Molecular Probes) or Alexa Fluor (AF)-594 succinimidyl ester (Molecular

Probes), was performed as described by Pelkmans et al. (2001). SV40, labelled with

Cy5, FITC or AF-594 dyes, was obtained from L. Pelkmans (Pelkmans et al., 2001).

2. Cell cultures (I-IV) and transfections (II, III)Cells. The cell lines used in the studies were African green monkey kidney cell lines

(CV-1, GMK), human epitheloid carcinoma cells (HeLa-Ohio) and human

osteosarcoma cells (HOS, SAOS-2). All the cell lines were obtained from ATCC.

HOS-pα2AW (Riikonen et al., 1995) and SAOS-α2β1 cells (Ivaska et al., 1999b)

were generated from wild-type cell lines that normally lack endogenous α2 integrin

by transfection of cells with a vector carrying the α2 integrin subunit (Ivaska et al.,

1999b). SAOS-α1/α2β1 cells expressed the chimeric form of α2 subunit in which the

cytoplasmic domain of α2 was replaced by that of α1 integrin subunit (Ivaska et al.,


49

1999b). SAOS-pAW cells, carrying the pAW vector, were used in control

experiments.

The cells were maintained in Dulbecco's MEM (DMEM; GibcoBRL), supplemented

with 10% foetal calf serum (FCS), 2 mM L-glutamine, 100 IU/ml penicillin, and 100

µg/ml streptomycin. Antibiotic G418 (200 µg/ml)(GibcoBRL) was added to

transfected SAOS and HOS cells. The infections were performed in the same media

but the concentration of FCS was reduced to 1% (infection media). Virus infections

were performed for 1 h on ice or alternatively for 1 h at 4°C to allow the binding of

the virus. The unbound virus was removed by washing and cells were transferred to

37°C.

Transfections. The transient transfections of CV-1 and/or SAOS-α2β1 cells were

performed with Fugene 6 (Roche) or Superfect (Qiagen). N- or C-terminally GFP-

tagged constructs of caveolin-1 (GFP-caveolin-1 and caveolin-1-GFP,

respectively)(Pelkmans et al., 2001) and HA-tagged caveolin-3 and caveolin-3DGV

(Roy et al., 1999) were used to investigate the role of caveolar route in infection.

GFP-tagged wt dynamin 2 and dominant negative dynamin 2(K44A) (Cao et al.,

1998; Ochoa et al., 2000), GFP-tagged wt Eps15 and Eps15E∆95/295-GFP

(Benmerah et al., 1999) and c-myc tagged adaptor protein 180 with truncated C-

terminus (AP180C construct)(Ford et al., 2001) were also used in transfections. The

exact functions of the constructs mentioned above are explained in the Results and

Discussion. In addition, a myc-tagged transferrin receptor construct was used to mark

the clathrin -dependent endocytosis route.

3. Infectivity and binding assays (I-IV)Plaque assay. The following plaque assay was used in several experiments: The

viruses were incubated with the cells on 6-well plates for 30 min at 37°C. After

removal of unbound virus, the cells were overlaid with 0.5 % carboxymethylcellulose

in infection media. After 2 days incubation at 37°C, the cells were stained with crystal

violet before counting of the plaques.

Infectivity titration. For determination of viral replication cycle, the cells were

infected with virus and collected at different time-points of infection. After three

freeze-thaw cycles, the amount of intracellular virus was determined by plaque assay.

Alternatively, infectivity titration in GMK cells was performed to harvested samples.

After incubation for 7 days at 37°C, the cells were stained with crystal violet for the

determination of end-point titres.

Inhibitory assays with drugs. For inhibitory assays, CV-1 cells were preincubated

with different cellular inhibitors for 30 min at 37°C. Purified EV1 was allowed to


50

bind to cells for 1 h at 4°C. After removal of the unbound virus, the cells were

incubated in the presence of the drugs for 6 h at 37°C. For testing the effects of

brefeldin A (BFA, Sigma), the CV-1 cells were incubated in the presence of the drug

(0.5-2 µg/ml) for 1 h at 37°C prior to the infection. Concentrations of the drugs used

were 5 µM bisindolylmaleimide (Sigma), 1-7 µg/ml of cytochalasin D (Sigma), 25-

250 µM genistein (Sigma), 0.13-1 µM latrunculin A (Molecular Probes), 0.1-0.75 µM

jasplakinolide (Molecular Probes), 10 mM methyl-β-cyclodextrin (Sigma), 33 µM

nocodazole (Sigma), 25 µg/ml nystatin (Sigma) together with 10 µg/ml progesterone

(Sigma), 1 µM okadaic acid (Sigma), 10 µM safingol (Sigma) and 1 mM sodium

orthovanadate (Calbiochem). The effects of the drugs on cellular functions are

explained in the original publication IV. After infection the cells were stained with

anti-EV1 antiserum for immunofluorescence microscopy or the infectivity of the virus

was determined by plaque assay.

Inhibition assay with the αααα2I domain. Plaque assay was used to test whether the

preincubation of viruses with their receptors could result in inhibition of the infection.

Prior to plaque assay, EV1 (200 plaque forming units; PFU) was incubated alone,

with different concentrations (0-1000 nM) of a recombinant glutathione-S-transferase

(GST) fusion protein of α2I (α 2I-GST) (obtained from J. Käpylä, University of

Jyväskylä, Finland) or with GST for 1 h at 37° C. Corresponding incubations were

performed for PV1 and PVR-Fc, obtained from L. Xing (Karolinska Institute,

Huddinge, Sweden) (Xing et al., 2000). The number of plaques obtained in the

presence of receptor molecules was compared to the amount of plaques observed in

the absence of receptor.

Antibody blocking assay and immunoperoxidase staining. The confluent SAOS-

α2β1 were preincubated with different antibodies or their combinations for 15 min at

RT: anti-α2 integrin (12F1; BD Pharmingen), anti-β2 microglobulin (polyclonal

rabbit antibody AB730; Chemicon and monoclonal antibody BM-63; Sigma) and anti

human HLA-I W6/32 (Barnstable et al., 1978). After 10 h incubation at 37°C with

EV1, the cells were fixed with methanol and stained with anti-EV1 rabbit antiserum

and horseradish peroxidase-conjugated swine anti-rabbit immunoglobulin (Dako,

Denmark) (Ziegler et al. 1988).

Cell binding assay. SAOS-pAW or SAOS-α2β1 cells, suspended into PBS

containing 1 mM MgCl2 (PBS-MgCl2) were incubated with [35S]-labelled EV1 for 1

h on ice. The cells were washed with PBS-MgCl2 and analysed for radioactivity in a

scintillation counter.


51

4. Sucrose gradient sedimentation (I-III)To study whether the binding of EV1 to GST-α2I domain can trigger conformational

changes in the viral capsid, [35S]-labelled EV1 was incubated with 100-1000 nM α2I-

GST for different time periods at 4°C and at 37°C. To investigate the uncoating of the

virus during the entry into host cells, [35S]-labelled virus was allowed to bind to

detached SAOS-α2β1 or GMK cells in suspension for 1 h at 4°C prior to incubation

of 30 min-2 h at 37°C. The cells were lysed with 1% Triton-X-100 for 10 min on ice.

The collected samples were centrifuged in a linear 5-20 % sucrose gradient and the

radioactivity of collected fractions was measured in a scintillation counter.

5. Immunofluorescence microscopy (I-III)After the binding of viruses to the cells in cold for 1 h and the infection at 37°C for an

appropriate time, the cells were fixed with 4% formaldehyde or 4% paraformaldehyde

(Sigma), quenched with 50 mM NH4Cl and permeabilized with 0.05% (w/v) saponin

(Sigma) or 0.3% Triton-X-100 (Sigma) in PBS. The primary antibodies described in

Table 3 were used for immunofluorescent labelling (Table 3). To recognize the

primary antibodies, we used AF-488- and 568–conjugated anti-mouse, anti-goat and

anti-rabbit secondary antibodies (Molecular Probes). Mouse anti-HA antibody

(BabCO) was used to recognize the HA-tag.

AF-488-conjugated cholera toxin subunit B (0.5 or 10 µg/ml; Molecular Probes) was

used in the EV1 internalization studies as a marker of the caveolar route. Lysotracker

Red DND99 (100 nM) (Molecular Probes) was used to mark the lysosomes and holo-

transferrin (1 mg/ml; Sigma) to detect the clathrin route. The labelled molecules were

either preincubated with cells prior to virus infection or added simultaneously with

the virus.

After staining with primary and secondary antibodies, the cells were mounted for

microscopy. Microscopy was performed either with an Axiovert confocal microscope

(Leica TCS SP2) with an HCX PL APO 63x/1.32-0.6 oil objective or with Axiovert

100 M SP epifluorescence microscope (Carl Zeiss, Jena, Germany) equipped with a

confocal setup (Zeiss LSM510), using a Plan Neofluar objective (63x/1.25 oil). For

quantification of infection percentages, images of 100-200 cells were taken with an

Olympus BX50 immunofluorescence microscope containing a CCD camera

(Hamamatsu), using a U Plan FI 20X/0.5 Ph1 objective and the Open Lab 2.2.5

program (Improvision). Further analysis of the results is explained in the

corresponding articles (I-III).


52

6. Real-time fluorescence microscopy (III)After binding of AF-EV1 to untransfected or GFP-tagged caveolin-1-transfected CV-

1 cells for 1 h at 4°C, the microscopy was performed at 37°C using a Zeiss Axiovert

wide-field microscope with a 100 X NA 1.4 plan-apochromat lens, a computer-

controlled shutter and standard FITC/Alexa Fluor-594 filters. Images were collected

with a CCD camera, with 2×binning, delay times of 4-10 s and exposure times of 0.5-

1 s/image. The collected images were further processed using Open Lab 2.2.5

software.

Table 3. Primary antibodies used for immunofluorescence microscopy.Target ofdetection

Antibody Manufacturer/Reference

Capsid proteinsof EV1

- polyclonal rabbit antiserumagainst EV1

(Marjomäki et al., 2002)

integrin αααα2222subunit

- MAb 12F1- MAb 1950- AF-488-labelled mouse antibodyMCA2025- rabbit antiserum

BD PharmingenChemiconSerotec

Transduction laboratoriesintegrin αααα2Idomain

- anti-CD49b monoclonal antibody Immunotech

Caveolin-1 - polyclonal rabbit antibody- rabbit antibody N-20- MAb 2234- MAb

Transduction LaboratoriesSanta Cruz BiotechnologyTransduction LaboratoriesZymed

Myc-taggedtransferrinreceptor chimera

- MAb against myc-peptide ATCC

Clathrin route - rabbit anti-transferrin antibody Behring InstituteEarly endosomes - polyclonal antiserum against

EEA1(Mu et al., 1995)

Late endosomes - polyclonal antiserum against thecation-independent mannose-6-phosphate receptor (CI-MPR)

(Marjomäki et al., 1990)

Lysosomes - MAb for CD63 ZymedTrans-Golginetwork

- MAb anti-p230- polyclonal rabbit antiserum GB2- TGN-46

Transduction Laboratories

(Banting et al., 1998)Cis-Golginetwork

- polyclonal antiserum against p23 (Rojo et al., 1997)

ER - MAb against PDI (ID3) from S. Fuller, University ofOxford, United Kingdom

sER - goat antibody against syntaxin 17 (Steegmaier et al., 1998)


53

7. Fluorescence in situ hybridisation (III)The negative-polarity RNA strand was transcribed using SP6 polymerase (Promega)

from linearized EV1 cDNA (full-length EV1 cDNA in the pSPORT1 vector) (Ohman

et al., 2001) in the presence of FITC-labelled UTP (Molecular Probes) or Chromatide

Alexa Fluor-546-14-UTP (Molecular Probes). After DNase I treatment, the probe was

purified as described earlier (Bolten et al., 1998). To perform fluorescence in situ

hybridisation (FISH), the untransfected and GFP-caveolin-1 -transfected CV-1 cells

were infected with EV1 and the hybridisation reaction was performed in the presence

of the labelled probe for 12 h at 42°C in the dark (Bolten et al., 1998). After removal

of the unbound probe, cells were mounted with 2.5% DABCO (Sigma) in pH-

buffered glycerol and visualized using a confocal microscope (Leica) as described

above.

8. Protein synthesis assay (IV)For the analysis of host cell protein synthesis during infection, the cells were infected

with purified viruses (EV1, CBV4, PV1) and incubated at 37°C until most of the cells

showed CPE. After starving of cells in the presence of MEM-met (Gibco) for 30 min,

they were labelled with 50 µCi/ml of [35S]methionine (Amersham). The cells were

either harvested directly after a pulse labeling for 15 min or chased in complete

medium (MEM) for additional 30 min at 37°C. The radioactively labelled proteins

were analysed in 12% SDS-PAGE.

9. cDNA array technique (IV)To study the effects of enterovirus infection on cellular gene expression, HOS

pα2AW cells were infected with purified EV1 (MOI 130 corresponding to 0.5 µg of

purified virus/1x106 cells). HeLa-Ohio cells were infected either with purified CBV4

or PV1 (MOI 20). The infected and mock-treated control cells were collected after

different time intervals. Total cellular RNA was purified using the thiocyanate-CsCl

method (Chirgwin et al., 1979). Five µg of total RNA was converted to specific 32P-

labeled cDNA species by using MMLV reverse transcriptase (Clontech), in the

presence of CDS primer mix (Clontech) and 10 mCi/ml of [α 32P]dATP (Amersham).

Equal amounts of 32P-labeled cDNA as cpm values from infected and uninfected cells

were hybridised to two identical filters, using both Atlas human cDNA array and

Atlas human cDNA 1.2 array (Clontech) according to the manufacturer’s instructions.

The filters were exposed on BioMAX MS film (Kodak) and phosphoimager screen

(Fuji). The photo-stimulated luminescence (PSL) -signals were measured by using

phosphoimager MacBas -quantitation program (Fuji).

Results and Discussion

54

RESULTS AND DISCUSSION

1. Cell surface interactions of echovirus 1 (I-III)

1111....1111 αααα2ββββ1 integrin as EV1 receptor (I-III)

To study the attachment of EV1 to the cell surface, the radioactively labelled

virus was allowed to bind to control SAOS-pAW cells, lacking the integrin

α2 subunit and to SAOS-α2β1 cells, which have been transfected with a

vector carrying the α2 subunit (Ivaska et al., 1999b). The binding assay

revealed that over 70% of virus particles were able to bind to SAOS-α2β1

cells whereas only 0.5% of particles were bound to SAOS-pAW cells (II, Fig.

2B; The figures referring to original publications are marked in italics). Only

SAOS-α2β1 cells were susceptible to EV1 infection, which was efficiently

inhibited (86%) by antibodies against α2 integrin in a immunoperoxidase

staining assay (II, Fig. 1 and 2A, C, D). In confocal studies of the infected

cells, the virus and the integrin colocalized on the cell surface immediately

after virus attachment on ice (0 h p.i.) (II, Fig. 4A). The α2β1 integrin also

colocalized with EV1 in the perinuclear accumulations after incubation at

37°C (II, Fig. 4A, III, Fig. 2C), indicating that the integrin plays also a role in

the internalization of the virus. In addition, the preincubation of EV1 with

integrin α2I domain-GST (α2I-GST) prior to addition of the virus to cells

blocked infection in a dose-dependent manner (I, Fig. 1A).

Overall, our results, in accordance to earlier reports (Bergelson et al., 1992),

showed that α2β1 integrin may be sufficient for EV1 attachment and cell

entry. However, the role of accessory receptors in EV1 entry cannot yet be

excluded. In our investigations, the antibodies against β2 microglobulin

(β2m) caused an efficient inhibition (95%) of EV1 infection in SAOS-α2β1

cells (II, Fig. 2D). Although β2m is coexpressed with MHC class I molecule

on the cell surface, the antibodies tested against MHC class I did not inhibit

infection. In immunofluorescence confocal microscopy studies, β2m was not

internalized into cells together with the virus and the integrin (II, Fig. 4B).


55

Because antibodies against β2m have been reported to block the infection of

several enteroviruses (Ward et al., 1998), the molecule may be unspecifically

involved in the virus life cycle. Alternatively, antibodies against β2m may

cause a steric hindrance to viral binding to other receptors.

1.2 The αααα2I domain binds to the EV1 canyon (I)The tools for structural studies of EV1-α2I domain interaction became

available when the structure of α2I domain was solved in 1997 (Emsley et al.,

1997), and the atomic structure of EV1 was visualised by x-ray

crystallography in 1998 (Filman et al., 1998). In our study, cryo-EM

reconstruction was used to illustrate the structural interactions of EV1 with

α2I-GST fusion protein (I).

Figure 9. A) The α2I domain (gray; fitted into the difference density map) interactswith two adjacent protomers of EV1 capsid. 5-fold and 3-fold axes are indicated. B)The charged residues important for EV1-α2I domain interaction, including Lys201,Asp219 and Arg288 in the I domain and Glu2162, Asp2163, Lys3230 and Glu1273 inthe virus are indicated. (Adapted from the original publication IV, Fig. 2 C, D).


56

Residues involved in αααα2222ΙΙΙΙ domain-EV1 interaction. The reconstruction of

EV1-α2I domain revealed that the virus is decorated by 60 copies of α2I

domain (I, Fig. 2B). α2I located roughly between the viral 2-fold and 5-fold

axis. The unliganded α2I domain structure (Emsley et al., 1997) was fitted

into the difference density map, generated by subtracting the crystal structure

based density of native EV1 (I, Fig. 2A) from the cryo-EM density map of the

virus-receptor complex (I, Fig. 2B). In the resulting model, both the N- and C-

termini of the I domain pointed outwards from the virus-interacting face (Fig.

9A), thus resembling the natural situation where these termini ends are

connected with a β-propeller structure of the α2-subunit.

The binding site of the α2I domain in the EV1 surface is situated mostly on

the outer (=south) wall of the viral canyon (Fig. 9A). Instead, the I domain is

not in intimate contact with the inner (=north) canyon wall. The α3 helix of

the α2I domain together with the connecting loops interact with the VP2 from

one protomer and the globular head contacts VP3 from a neighbouring

protomer of EV1 (Table 4, Fig. 9A).

Table 4. The interacting residues in α2I domain and EV1. The viral proteins andresidues involved in the interactions have been resolved in the cryo-EM study of α2I -EV1 complex (I).

STUDY METHOD αααα2I RESIDUES EV1 PROTEINS EV1 RESIDUES

- Cryo-EM reconstruction

- Chimeras of human and

mouse α2I (King et al.,

1997)

- Tyr200, Lys201

- 199-201between C-strandand α helix 3

E-F loop of VP2Glu2162Asp2163His2164

- Cryo-EM reconstruction

- Chimeras of human and

mouse α2I (King et al.,

1997)

- Tyr216- 212-216bw. α helix 3 and

α helix 4

above the interfaceof two adjacentprotomers

Tyr216 of α2Ipoints toward theviral surface

- Cryo-EM reconstruction Asp219 VP3 Lys3230

- Cryo-EM reconstruction Arg288 in

αC-α6 loop

the C-terminus ofVP1 of the 2ndprotomer

Glu1273

- Chimeras of human α1I

and human α2I (Dickeson

et al., 1999)

Arg289 in αC-α6

loop

the C-terminus ofVP1 of the 2ndprotomer

No direct contact,conformationaleffect possible


57

The MIDAS points towards the canyon floor but it is not in close contact with

the virus. The charged residues (Lys201, Asp219 and Arg288) of α2I domain

interact most probably with viral residues Glu2162-Asp2163, Lys3230, and

Glu1273 (numbered to start as 1001 for VP1, 2001 for VP2, 3001 for VP3

and 4001 for VP4) (Fig. 9B). The earlier studies with a mutated integrin α2

subunit and with integrin chimeras have shown that the amino acids in regions

199-201 and 212-216 (King et al., 1997) as well as Arg289 (Dickeson et al.,

1999) in α2I domain interact with the virus. Most of the corresponding

residues are in contact with the virus according to our cryo-EM reconstruction

of EV1-α2I domain complex, as shown in Table 4.

Like α2β1 integrin, the IgSF receptors of major group HRVs, PVs, and CBV3

bind to the viral canyon (He et al., 2001). In contrast, non-IgSF receptors, like

LDLR (minor group of HRVs) and DAF (EV7) bind outside the canyon

(Hewat et al., 2000; He et al., 2002). When compared to PVR-PV interaction

(Xing et al., 2000), both ICAM-1 (Olson et al., 1993; Xing et al., 2000) and

the α2I domain are oriented deeper towards the canyon floor and the pocket

factor cavity of the viruses. Both ICAM-1 (Olson et al., 1993) and the α2I

domain primarily interact with the outer wall of the canyon. In contrast, the

PVR has been argued to contact either both the outer and inner walls (Belnap

et al., 2000b; He et al., 2003) or predominantly the outer wall of PV (Xing et

al., 2000). Whatever the case, the α2I domain makes more extensive contacts

with the outer canyon wall than ICAM-1 or PVR. Moreover, compared to

footprints of ICAM-1/PVR on the major group HRVs/PVs, the footprint of

the α2I domain on EV1 is more globular.

Structural data on other picornavirus-integrin interactions are not yet

available. These interactions may anyhow differ significantly from EV1-α2β1

integrin interactions, since they occur through the viral RGD sequence (Chang

et al., 1989; Fox et al., 1989; Hyypiä et al., 1992), not present in EV1 capsid.

Moreover, αv integrins do not carry the αI domain. Instead, the I-like domain

of β-subunit is involved in the integrin interactions with the RGD motif

(Xiong et al., 2002).


58

1.3 The differences in interactions of αααα2ββββ1 integrin with EV1 andcollagen (I)

In our study, a crystal structure of collagen-like triple helical peptide bound

on the α2I domain (Emsley et al., 2000) was superimposed into the model of

EV1-α2I domain (I, Fig. 3A, B). Extensive overlap between collagen and the

virus was observed, indicating that these two ligands could not bind the

integrin simultaneously even though their binding sites in the α2I domain are

distinct, as also shown in other studies (Kamata et al., 1994; King et al.,

1997). The MIDAS site, known to interact with the middle strand of collagen

(Emsley et al., 1997; Emsley et al., 2000), is not involved in EV1-α2β1

interaction as demonstrated here and elsewhere (Bergelson et al., 1993a; King

et al., 1997). In the cryo-EM model of EV1-α2I, the MIDAS site points

towards the canyon floor of EV1 and thus it is inaccessible for simultaneous

collagen binding. However, there is enough space to simultaneously

accommodate another MIDAS-binding element, a cyclic RKKH -containing

octapeptide (Ivaska et al., 1999a). The peptide has been shown to block

collagen binding to the α 2I domain but to increase EV1 binding, most

probably by increasing the contact surface between the I domain and virus

and by shielding the negatively charged residues of the MIDAS.

Adhesion of collagen to α2β1 can be abolished by chimeras containing the

cytoplasmic domains of other integrin subunits in place of α2 tail (Kawaguchi

and Hemler, 1993). However, the virus could infect chimeric SAOS-α2/α1β1

cells (II, Fig. 2). Therefore, the α2 cytoplasmic domain is not required for

EV1 entry or infection, in line with the earlier results (Bergelson et al.,

1993a). Further studies should be performed to find out whether the

cytoplasmic domain of β1 integrin subunit, instead of the α2 tail, could play a

role in signalling events related to EV1 endocytosis (Upla et al., 2004).

The buried surface area of I domain was much larger (850Å) when bound to

EV1 compared with collagen-like peptide (359Å), indicating that the affinity

of virus to the I domain could be higher than that of collagen (I). The binding

affinity of EV1 to the I domain, measured by solid phase binding assay, was

found to be about 10 times higher when compared with type I collagen (I,

Fig. 3C). Thus, in tissue, the virus has either to compete with collagen on


59

binding to α2β1 integrin or find ligand-free integrin molecules. Normally,

α2β1 integrin is expressed on basolateral surface of cells, where it is occupied

by collagen or other natural ligands. However, in some cell types, such as in

keratinocytes, the integrin may be expressed on free from natural ligands

(Larjava et al., 1993) and thus it may be more susceptible to virus infection.

1.4 EV1 binding to αααα2ββββ1 integrin may trigger integrin clustering (I)

The confocal studies of α2β1 integrin showed the formation of integrin

clusters on the cell surface after the virus attachment and a similar clustering

was induced by cross-linking the integrin molecules with antibodies (I, Fig.

4). To study whether EV1 could bind many integrin molecules

simultaneously, a model of α2β1 heterodimer was constructed based on the

crystal structures of extracellular parts of αvβ3 (Xiong et al., 2001) and α2I

domain (Emsley et al., 1997). The model was then superimposed on α2I

domain - EV1 structure. As a result, five copies of the integrin molecule were

found to bind without steric hindrance around the viral five-fold axis, possibly

resulting in integrin clustering (I, Fig. 5). This is in accordance with other

findings suggesting that the attachment of EV1 to α2β1 integrin can induce

the clustering of integrin molecules, which in turn leads to signalling events,

required for endocytosis of EV1 (III, Upla et al., 2004). Interestingly, binding

of adenovirus to the αvβ5 integrin may lead to similar clustering of integrin

molecules (Chiu et al., 1999), which is followed by signalling events essential

for virus entry into host cells (Greber, 2002).

1.5 The uncoating of EV1 is not triggered by cell surfaceinteractions (I-III)

To identify whether the cell surface interactions of EV1 are able to trigger the

uncoating of the virus, EV1 was incubated alone, with the α2I-GST or with

GMK or SAOS-α2β1 cells and the conformational alterations of the virus

were measured by sucrose gradient centrifugation. At 4°C, the viral capsid

was observed in 160S form of intact virus that carries the genome (I, Fig. 1B).

Incubation of EV1 at 37°C for 1 h induced a spontaneous uncoating of the


60

virus, leading to the formation of 80S empty particles (I, Fig. 1B). However,

the incubation of EV1 with α2I-GST at 4°C, 20°C and 37°C for different time

periods caused only minor changes in virus sedimentation and did not lead to

formation of 135S or 80S particles (I, Fig. 1B).

It could be speculated that the binding of the entire α2β1 to EV could bring

about different actions other than the α2I-GST alone or that some other cell

surface molecules could induce the conformational changes of EV1 particle.

However, only minor alterations in virus sedimentation, somewhat similar to

those observed in the presence of the α 2I domain, were observed after

binding of EV1 onto cell surface at 4°C (II, Fig. 3A, III, Fig. 8A). When

compared to control 135S particles of PV, the altered EV1 particles

sedimented again much closer to the 160S peak. The studies with mutant PVs

have revealed another intermediate of viral entry, the 147S particle, which

may sediment close to 160S particle and which may be much less stable than

135S (Pelletier et al., 2003). Therefore, further investigations are required to

clarify the nature of altered EV1 particles. On the other hand, the minor

changes observed in virus sedimentation could be due to the formation of the

EV1-α2I domain/integrin complex.

Our results suggest that cell surface interactions of EV1 do not lead to the

instant uncoating of the virus on the cell membrane, even though the slight

alterations may be caused in the virion. Moreover, the α2I domain may

stabilize EV1 structure in vitro, or at least inhibit the formation of 80S empty

particles. This is different from many other picornavirus-receptor interactions,

such as PV-PVR and HRV-ICAM-1, which lead to obvious conformational

changes in the viral capsid and formation of 135S particles. Finally, these

interactions may result in the formation of 80S particles and release of

genomic RNA during the incubation at 37°C (Xing et al., 2000). The

stabilization of EV1 capsid by α2I domain might ensure that the virus is

endocytosed to the location adjacent to the replication site before the release

of viral genome is triggered by intracellular factors.


61

2. The endocytosis of echovirus 1 into caveosomes (II, III)

Because integrins are known to mediate the internalization of several

pathogens, it was of interest, whether the α2β1 integrin could guide EV1 into

a specific endocytic route (II, III). To study the uptake of EV1 into host cells,

both microscopic methods (e.g. EM, immunofluorescence microscopy and

real-time microscopy) and biochemical and virological approaches (e.g.

fluorescence in situ hybridisation (FISH), infectivity assays and sucrose

gradient centrifugation) were applied. SAOS-α2β1 cells were used in studies

of EV1 endocytosis because they overexpress the α2β1 integrin. Thus, the

visualization of the integrin in this particular cell line is facilitated in

fluorescence microscopy. CV-1 cells were primarily chosen because of their

excellence in real-time live microscopy studies, and because the caveolar

endocytosis of SV40 has been well-characterized in this particular cell line.

The infectious cycle of EV1, detected by infection titration, took

approximately 6-8 h in these cell lines (II, Fig. 2E, III, Fig. 1B). The genomic

viral RNA, detected by FISH technique, was observed to increase at 4 h p.i. in

CV-1 cells (III, Fig. 8C). The increase indicated the synthesis of new viral

RNA and thus suggested that viral replication takes place at 3-4 h p.i.

2.1 EV1 does not utilize clathrin-mediated endocytosis (II, III)

In double-labelling immunofluorescence microscopy, EV1 colocalized with

the α2β1 integrin on the cell surface (II, Fig. 4). Soon after binding, already

at 5 min p.i., the virus started to internalize together with the integrin into the

host cells. Even though receptor-mediated uptake is a typical characteristic of

clathrin-mediated endocytosis, the virus or its receptor were not found in

organelles of the clathrin route, such as EEs, recycling endosomes or LEs by

immunofluorescence microscopy (II, Fig. 5). When the formation of clathrin-

coated vesicles was prevented by expression of Eps15E∆95/295-GFP

(Benmerah et al., 1999), no inhibition of EV1 infection was observed in CV-1

or SAOS-α2β1 cells (III). Overexpression of AP180C, which interferes with

the clathrin route and uptake of HRV2 (Ford et al., 2001; Snyers et al., 2003)


62

did not exert this inhibitory effect either. Accordingly, the same constructs

have revealed no effect on the internalization of the α2β1 integrin after

antibody-crosslinking (Upla et al., 2004).

The clathrin route of entry is utilized among picornaviruses, for instance, by

minor group HRVs, which take advantage of endosomal acidification for

uncoating (Bayer et al., 1998). Instead, enteroviruses, like EV1, are

distinguished from other picornaviruses by their acid stability and may thus

have different requirements for the milieu of uncoating. Moreover, the

specific location of viral receptor on the cell surface may influence the

selection of endocytosis route. For example, acid-stable EV11 binds to a

receptor (DAF) located in lipid rafts, and then enters the cells via lipid

raft/caveolae-mediated endocytosis (Stuart et al., 2002a). Interestingly,

integrins can also be associated with lipid rafts (Upla et al., 2004) and

caveolae (Wary et al., 1998).

2.2 EV1 may utilize both cell surface caveolae and an alternativepathway to enter host cells (II, III)

Endocytosis via caveolae. The EM of EV1-infected SAOS-α2β1 cells

demonstrated the localization of the virus in caveolae-like invaginations on

the cell surface (II, Fig. 6B). More detailed studies on location of the viral

receptor, α2β1 integrin, have suggested that the integrin is first associated

with raft-like membrane microdomains, from where it is laterally distributed

to cell surface caveolae upon EV1 binding (Upla et al., 2004). In fact, both αand β subunits of α2β1 integrin can also associate with caveolin-1, the main

protein component of caveolae (Wei et al., 1996; Wary et al., 1998).

Several known inhibitors of caveolar internalization and dominant negative

mutants of the pathway were able to prevent EV1 uptake and/or infection (II,

Table 1, III, Fig. 3). Expression of dominant negative caveolin-3DGV, which

inhibits SV40 infectivity by 40% (Roy et al., 1999), diminished EV1 infection

by 35% in CV1 cells and by 66% in SAOS-α2β1 cells. Caveolin-3DGV

disrupts lipid transport and causes depletion of lipid rafts, thus inhibiting

caveolar endocytosis (Roy et al., 1999). The dominant-negative mutant of


63

dynamin 2 (K44A), which prevents the caveolar and clathrin-coated pit

internalization (Damke et al., 1994; Henley et al., 1998; Oh et al., 1998),

inhibited EV1 infection by 75% in CV-1 cells. Moreover, drugs interfering

with the composition of lipid rafts and caveolae and with the cholesterol

distribution (methyl β-cyclodextrin, progesterone and nystatin) hindered EV1

infection. In addition, methyl β-cyclodextrin decreased the intracellular

colocalization of the virus, caveolin-1 and the integrin by over 90% in SAOS-

α2β1 cells.

Tyrosine kinase activation is essential for the EV1 replication cycle since the

infection was blocked by a tyrosine kinase inhibitor, genistein, in a dose-

dependent manner in CV-1 cells. Genistein is known to inhibit the caveolar

uptake of several other ligands, including AMF, CTX and SV40 (Dangoria et

al., 1996; Le and Nabi, 2003). Furthermore, bisindolylmaleimide, a specific

inhibitor of protein kinase Cα (PKCα), prevented EV1 infection in CV-1 cells

(III, Fig.6). PKCα activation has also been demonstrated in EV1-infected

SAOS-α2β1 cells, where it triggers viral entry and leads to activation of ERK

(Upla et al., 2004). Interestingly, PKCα is known to bind to the cytoplasmic

domain of the β1 integrin subunit and to regulate cell motility (Ng et al.,

1999). Our results are also in line with the fact that caveolae-enriched PKCα(Mineo et al., 1998) is required for stimulation of caveolar internalization

(Sharma et al., 2004).

Taken together, the results suggested that EV1 particles may utilize dynamin-

dependent uptake via cell surface caveolae and that the uptake requires

particular signalling events.

An alternative route of endocytosis. A number of findings suggested that

EV1 might, in addition to uptake via cell surface caveolae, use an alternative

pathway for endocytosis. When endocytosis of fluorescently labelled EV1

was followed into GFP-caveolin-1 transfected CV-1 cells by real-time live

microscopy, the majority of EV1 particles did not colocalize with GFP-

caveolin-1 at the very beginning of infection (III). Dominant-negative GFP-

construct of caveolin-1, in which the N-terminal domain is truncated


64

(caveolin-1-GFP), did not inhibit EV1 internalization or infectivity in CV-1 or

SAOS-α2β1 cells (III), even though it has been reported to prevent the

uptake of SV40 through caveolae (Pelkmans et al., 2001). Our results

indicated that either the N-terminus of caveolin-1 is not required for

interactions EV1 in cell surface caveolae or the virus utilizes a different

uptake mechanism than SV40 in CV-1 cells.

For further evidence of an alternative pathway, some inhibitors, described to

affect caveolar internalization, had unexpected effects on the EV1 infectious

cycle (III, Fig. 6B). Such inhibitors included okadaic acid (an inhibitor of

serine and threonine phosphatases), which should increase caveolar

internalization (Parton et al., 1994) and sodium orthovanadate (an inhibitor of

protein tyrosine phosphatases) that enhances the uptake of SV40 via caveolae

(Pelkmans et al., 2002). Unexpectedly, okadaic acid did not significantly

affect EV1 infection and sodium orthovanadate reduced the viral infectivity

by 70%. However, the effects of these drugs were tested on EV1 infectivity

and not specifically on virus internalization. Therefore, they may also disturb

later steps of EV1 infection. The actin-disturbing agents, such as latrunculin

A, cytochalasin D and jasplakinolide, did not interfere with EV1 infection in

CV-1 cells (III, Fig. 6C), again opposite from the fact that internalization of

ligands through cell surface caveolae requires cortical actin (Parton et al.,

1994; Pelkmans et al., 2002). In contrast, in SAOS-α2β1 cells, the actin

polymer-stabilizing drug jasplakinolide caused the maintenance of virus and

the integrin on the cell membrane and inhibited infection.

The results suggested that depending on the cell line and cellular

environment, EV1 can switch from caveolar pathway to another

internalization pathway. This could occur, for instance, when the function of

plasmamembrane caveolae is inhibited. This hypothesis is in line with recent

studies, which have revealed that alternative pathways of entry are utilized by

other viruses and bacterial toxins (Damm et al., 2005; Kirkham et al., 2005).

Whilst our findings favour the model that the non-caveolar cell surface lipid

rafts are involved in the uptake of EV1 via an alternative pathway, further

investigations are required to establish their role in EV1 entry.


65

2.3 EV1 is internalized into caveosomes (II, III)

The immunolabelling of cryosections from EV1-infected cells showed the

colocalization of the virus and caveolin-1 in uncoated, intracellular vesicles of

60-90 nm (II, Fig. 6C, E). These structures were immunoisolated from the

infected cells with beads coated with caveolin-1 antibody that recognizes

caveolin-1 only in intact vesicle structures (Oh and Schnitzer, 1999) (II, Fig.

8). Based on immunisolation, the virus and α2β1 integrin were detected in the

caveolin-1 -positive fractions in significantly increased amounts at 15 min p.i.

Live microscopy of CV-1 cells, transfected with GFP-tagged caveolin-1 and

infected with the fluorescently labelled EV1, revealed a dramatic increase in

the colocalization of the virus and caveolin-1 after 10 min p.i. (III).

Approximately at 20-30 min p.i., the larger clusters of caveolar vesicles were

observed in the electron micrographs of EV1-infected SAOS-α2β1 cells (II)

and in the real-time microscopy of CV-1 cells (III). The majority of these

organelles were mobile, and they preformed random, short (3-6 µm) or

medium distance (9-14 µm) movements with velocities of 0.02-0.3 µm/s. A

continuous sorting of virus particles between the vesicles was observed. The

behaviour and the outlook of the vesicles detected with live microscopy were

similar to caveosomes, organelles involved in SV40 uptake (Pelkmans et al.,

2001) and cholera toxin (CTX) (Montesano et al., 1982; Nichols, 2002).

Therefore, the colocalization of EV1 with these markers of caveosomal route

was investigated.

EV1 colocalizes partially with CTX and SV40 in caveosomes.

Fluorescently labelled CTX colocalized partially with EV1 at 30-60 min p.i.

in intracellular vesicular structures, distributed throughout the cytoplasm.

After 1 h p.i., CTX had accumulated into the Golgi complex, without an

apparent colocalization with EV1 capsid proteins (III, Fig. 4A). However,

CTX is not an accurate marker of caveosomes, because it can also traffic from

caveolar vesicles directly into EEs of the clathrin-coated pathway (Pelkmans

et al., 2004), or it may even enter via non-clathrin, non-caveolar endocytosis

(Kirkham et al., 2005). In the light of these reports, further studies are

required to specify the organelles where EV1 and CTX meet. However, based


66

on our other findings listed here, the organelles seem to represent

caveosomes.

The entry of SV40 and EV1 to coinfected CV-1 cells was followed in real-

time live microscopy and by immunofluorescence confocal microscopy (III,

Fig. 5A, B). The partial colocalization of EV1 with SV40 in small, fairly

immobile structures was observed at 15-30 min after EV1 entry. Upon

infection, the colocalization of the viruses as well as the size and the mobility

of organelles, where they colocalized, increased. To artificially accumulate

the majority of SV40 particles into caveosomes, CV-1 cells were treated with

nocodazole, a microtubule-disrupting agent that does not affect the uptake of

SV40 into caveosomes but inhibits its traffic to the ER (Pelkmans et al., 2001)

(III, Fig. 5C). The drug treatment resulted in an increased colocalization of

SV40 and EV1, again supporting the hypothesis that EV1 is transported into

caveosomes.

Brefeldin A (BFA), a widely used inhibitor that disrupts the Golgi and also

inhibits the different steps of caveolar endocytosis of SV40 and polyomavirus

(Norkin et al., 2002; Richards et al., 2002) inhibited EV1 infection most

effectively when added before 3 h p.i. (III, Fig. 7A). In the presence of BFA,

both EV1 and SV40 accumulated in coinfected cells into large, caveolin-1 -

positive organelles (III, Fig. 7B) that showed no staining with endosomal (CI-

MPR) or lysosomal (CD63) markers. Based on our results, the drug may

cause the accumulation of EV1 into caveosomes by preventing the uncoating

of EV1, or the release of EV1 from caveosomes by stabilizing the caveosomal

membranes. Moreover, BFA may inhibit the viral RNA replication, as in the

case of another enterovirus, PV (Maynell et al., 1992).

These findings indicate that the caveolin-1 -positive structures in which EV1

was transported from the cell surface together with its receptor, α2β1 integrin,

are caveosomes. The uptake of both SV40 (Pelkmans et al., 2001; Pelkmans

et al., 2002; Damm et al., 2005) and EV1 into caveosomes appears to be

caveolae/non-caveolar lipid raft -derived, tyrosine kinase -dependent and

largely by-passes endosomes and lysosomes (Figure 10). However, the


67

detailed endocytic mechanisms of these viruses differ in many aspects,

highlighting distinct roles of caveosomal route in virus infections.

Figure 10. The proposed internalization routes of EV1 and SV40 into caveosomes. Inaddition to "classical" caveolae-mediated endocytosis, SV40 can enter thecaveosomes via more rapid alternative pathway, which is dependent on noncaveolarlipid rafts (Damm et al., 2005). (Modified from Pietiäinen et al., submitted).

2.4 EV1 remains in caveosomes prior to replication (III)

To investigate whether EV1 capsid proteins are transported to the ER or to the

Golgi complex, several antibodies were used to mark these organelles in

immunofluorescence microscopy of the infected cells (II, Fig. 5, III, Fig. 4B).

However, the viral proteins did not colocalize with any tested markers for the

smooth and rough ER or the trans- or cis-Golgi networks in SAOS-α2β1 cells

at 2 h p.i (II, III). In CV-1 cells, EV1 was not detected in the trans-Golgi nor

in the ER at 2-4 h p.i (III). Based on these results, we concluded that EV1

capsid proteins do not enter the ER or the Golgi complex, but may remain in


68

caveosomes. This is in contrast to SV40 and polyomavirus, which are carried

from caveosomes into the (s)ER in a microtubule-dependent manner

(Pelkmans et al., 2001; Gilbert and Benjamin, 2004). Instead, the traffic of

ligands from the cell surface to caveosomes does not always require

microtubules (Pelkmans and Helenius, 2003).

The transport of some caveosomal vesicles carrying fluorescently labelled

EV1 seemed very similar to microtubular transport when followed in real-

time microscopy. However, the disruption of microtubules by nocodazole did

not apparently affect EV1 distribution or infectivity, detected by

immunofluorescence confocal microscopy (III). Most probably, viral proteins

can utilize microtubules for trafficking within the host cell, but these are not

obligatory for the infectious cycle, since the virus may not need to be

transported further from caveosomes.

The sucrose gradient analysis of conformational changes of EV1, after

incubation of the radioactively labelled EV1 with SAOS-α2β1 and GMK

cells for 30 min-2 h at 37°C, suggested that disassembly of the virus may

occur after it has been endocytosed (II, Fig. 3B, III, Fig. 8B). This was in

accordance with the results obtained with FISH technique, where a probe

recognizing a positive-stranded, genomic viral RNA was hybridised into EV1

-infected, GFP-caveolin-1 -transfected CV-1 cells (III, Fig. 9B). At the

beginning of the infection, the viral RNA did not clearly colocalize with

caveolin-1. However, at the time of the uncoating (1-2 h p.i.) the viral RNA

colocalized mostly with EV1 capsid proteins in caveosomes, and the capsid

proteins were not found in any other cellular compartments at 1-4 h p.i.

These results may indicate that the virus resides within caveosomes until

uncoating and replication take place, the phenomenon not observed with any

other ligands of caveosomal route. Therefore, it would be of great interest to

investigate the release mechanisms of EV1 and/or viral RNA from

caveosomes.


69

3. Echovirus 1 infection results in alterations of host cell geneexpression (IV)

The previous studies on cellular effects of EV1 have focused on individual

genes and their activation by MAP kinase pathways (Huttunen et al., 1997;

Huttunen et al., 1998). The availability of novel cDNA array technology

encouraged us to undertake a broad screening of the host cell gene

transcription profile during EV1 infection (IV). cDNA copies of cellular

mRNAs were isolated from uninfected and infected cells and hybridised onto

cDNA arrays. The human cDNA arrays contained either 588 genes (assayed

for EV1 -infected cells at 1, 3, 6, and 10 h p.i.) or an additional 588 genes

(assayed for EV1 -infected cells at 10 h p.i.). HOS-pα2AW cell line was

selected for the experiments because the cell line has been used in previous

studies of EV1-induced host cell gene regulation (Huttunen et al., 1997;

Huttunen et al., 1998).

At earlier time points of EV1 infection (1, 3, 6 h p.i.), no significant

alterations (i.e. over 2-fold) of host cell gene expression were observed (IV).

At 10 h p.i., the virus production in HOS-pα2AW cells had reached its

maximum. When the samples from 10 h p.i were screened with cDNA arrays

containing 1176 genes, only 2% of sequences were found to have a significant

(2-fold) increase in expression. Meanwhile, less than 0.5% of investigated

host cell genes were downregulated during EV1 infection. Such a low amount

of downregulated genes was somewhat unexpected, because enteroviruses, at

least PVs, should be able to inhibit cellular RNA synthesis (Rubinstein and

Dasgupta, 1989). On the other hand, many cellular genes are preferentially

upregulated during virus infections (Jenner and Young, 2005). The products

of the activated genes were involved in cellular signalling events, apoptotic-

and antiapoptotic functions, cell-cell interactions, DNA replication and in

regulation of transcription of target genes (IV, table 1).

We also studied whether there exist any similarities in host cell gene

regulation during infection of different enteroviruses. For this purpose, HeLa

cells were infected with CBV4 or PV1 for 7 h and the host cell gene

alterations were studied with cDNA arrays containing 588 genes. Some of the


70

altered genes found in our study are discussed below in more detail (IV, Fig.

2, Table 1, Fig. 3).

3.1 The upregulation of immediate early genes

At 10 h p.i., over 2-fold expression of several IE genes, including Fos-related

antigen (fra-1), c-jun, and early-growth response protein 1 (egr-1), was

observed in EV1-infected cells (IV, table 1, Fig. 2B). Both Fra-1 and Egr-1

were also upregulated, but less than 2-fold, at 6 h p.i (IV, Fig. 2C). Fra-1 can

heterodimerise with c-jun to form stable AP-1 complexes (Cohen et al., 1989)

that further regulate the transcription of target genes, involved in wide range

of cellular processes. Egr-1 regulates the genes involved in cell growth,

immune response and apoptosis by transactivation of the p53 gene (Thiel and

Cibelli, 2002). Therefore, it is the common link between the diverse pathways

that cause tissue injury (Yan et al., 2000). In addition, upregulation (2.5-fold)

of a gene encoding a transforming growth factor-β-inducible early growth

response protein (TIEG), was detected using another cDNA array with

additional 1176 genes (our unpublished data). TIEG can induce apoptosis and

regulate cell proliferation (Tachibana et al., 1997). Upregulation of IE genes

c-jun, c-myc and pim-1 (IV, Fig. 3) and TIEG (Johannes et al., 1999) was also

observed during PV1 infection in HeLa cells. Accordingly, activation of IE

gene expression has been detected during CBV3 infection (McManus et al.,

2002). Viruses belonging to other families can also regulate the expression of

IE genes, suggesting that these genes are mainly involved in general rather

than specific events in host cells during viral infections (Jenner and Young,

2005).

The increased expression of IE genes c-fos, c-jun and junB during EV1

infection has been shown to occur via MAPKs pathways (Huttunen et al.,

1998). Additional studies are required to specify which MAPKs could activate

the overexpression of fra-1, egr-1 and TIEG in EV1 infection. The gene

encoding MAPK-activated protein kinase 3 (MAPKAPK3), which is an

integrative element of MAPKs signalling in both mitogen and stress

responses, was upregulated in both PV and CBV4 -infected HeLa cells (IV,


71

Fig. 3). The enhanced expression of the MAPKs may play a role in regulation

of apoptotic events in the host cell during enterovirus infection, as shown

previously for CBV3 (Huber, 1999).

3.2 The upregulation of IRES-containing cellular genes

Based on metabolic labelling of host cell and viral proteins during EV1

infection, the virus was able to cause a partial inhibition of host cell protein

synthesis (shut-off) in HOS-pα2AW cells in the presence of visible CPE (IV,

Fig. 1B). In contrast, both PV1 and CBV4 induced more complete shut-off

effect in HeLa cells. The results implicate that control of host cell protein

synthesis varies between closely related viruses and may also vary depending

on the cell type.

Those cellular genes, which contain an IRES element, can be efficiently

translated in the presence of enterovirus-induced shut-off (Hellen and Sarnow,

2001). In the presence of a complete shut-off effect in PV1-infected cells, the

upregulation of pim-1 and c-myc mRNAs was detected here (IV) similiar to

earlier findings (Johannes et al., 1999). Somewhat surprisingly, the c-myc

gene was the only currently known IRES-containing gene, which became

elevated during EV1 infection. On the other hand, also those host cell genes,

which lack an IRES element can be translated in EV1-infected cells because

the virus does not cause a complete inhibition of protein synthesis.

The IRES-containing mRNAs encode proteins that are essential for cell

survival during a variety of cell stress situations (Hellen and Sarnow, 2001).

Because the c-myc gene product is a transcription factor that can control cell

vitality and induce apoptosis (Dang, 1999), the overexpression of c-myc gene

during enterovirus infection could, for example, present such a host stress

response to infection.

3.3 Other upregulated genes

During EV1 infection, many genes related to cell survival and apoptosis

were upregulated. Among them were the apoptotic IE genes c-myc, egr-1 and

TIEG mentioned above, and a gene coding for an anti-apoptotic myeloid


72

leukaemia cell differentiation protein (MCL-1), which is degraded, for

example, in adenovirus-induced apoptosis (Cuconat i et al., 2003).

Interestingly, a recent cDNA array comparison of 77 different host-pathogen

interactions revealed that overexpression of MCL-1 is highly common in

pathogen-activated cells (Jenner and Young, 2005). MCL-1 is regulated

through ERK-1-mediated signalling pathway (Michels et al., 2005), which is

activated at 5 h after EV1 infection (Huttunen et al., 1998). However, further

studies on these genes are required to solve whether EV1 regulates the

balance between antiapoptotic and apoptotic pathways like PVs (Belov et al.,

2003).

The expression of several transcriptional and translational regulators,

e.g. nuclease sensitive element binding protein 1 (NSEP1; YB-1) and

ribosomal protein L6 gene, coding for a protein component of ribosomal 60S

subunit (Zaman, 1993), were increased in EV1 infection. NSEP1 regulates

DNA-dependent transcription, modulates viral gene expression of HIV

(Kohno et al., 2003), and facilitates adenovirus replication (Holm et al.,

2002). Here, the overexpression of these genes may enhance the transcription

and translation of cellular genes important for EV1 infection and/or for host

cell defence mechansims.

Cell stress -related genes, such as growth arrest and DNA-damage-inducible

protein (GADD45) and a cAMP-dependent transcription factor 4 (ATF4) were

highly overexpressed in EV1 infected cells at 10 h p.i. The activation of

Gadd45 gene is also observed in apoptosis (Sheikh et al., 2000), in DNA

repair (Smith and Seo, 2002) and in several virus infections (Jenner and

Young, 2005). Genes of the ATF family have also been observed to be

upregulated during CBV3 (Taylor et al., 2000) (McManus et al., 2002) and

PV1 infections (Johannes et al., 1999). Thus, the universal laws of stress

responses may also apply to viruses.

A gene encoding the monocyte chemotactic protein 1 precursor (MCP-1), a

chemokine, was 2.9-fold upregulated at 10 h of EV1 infection. MCP-1 has

also been found to be upregulated following picornavirus (CVB3 and CVB4)


73

infection in human astrocytes (Kwon et al., 2004), after influenza A virus

infection (Sprenger et al., 1996) and in HIV-1 infection in mice (Potash et al.,

2005). Therefore, the induction of MCP-1 most probably represents a general

immune reaction against pathogens.

In summary, the overexpression of IE genes, stress response genes and genes

related to cell survival and apoptosis appears to be related to viral infections

in general. In contrast, upregulation of the cellular IRES-containing genes in

the presence of cellular protein synthesis shut-off may represent a specific

host cell response to those viruses, which interfere with cap-dependent

translation. In future studies, it would be interesting to find out whether the

induction of genes coding for certain kinases (e.g. CDC-like kinase 1, a

threonine-tyrosine kinase; IV, Table 1) could be particularly important in the

conversation between EV1 and the host cell. Moreover, further studies with

other techniques such as Western blotting are required to confirm whether the

upregulated genes are translated into functional proteins in the infected cells.

So far, the discussion about the functions of (up)regulated genes and their

products in EV1 infection remains rather speculative. Nevertheless, the study

provides significant data for further investigations of EV1-mediated signalling

events and viral pathogenesis.

Conclusions

74

CONCLUSIONS

The articles I-IV included in the thesis illuminate cellular interactions of

echovirus 1, a human pathogen. The basic mechanisms of virus - host cell

interactions share common properties amongst all viral pathogens. However,

the detailed contacts between viruses and host cells can vary within the

closely related viruses and even be serotype-specific, as illustrated here with

EV1.

The cryo-EM reconstruction of EV1 in complex with the α2I domain of its

receptor, α2β1 integrin, provides the first structure of integrin bound to

picornavirus. The integrin binding site in EV1 canyon is closely similar to the

binding sites of ICAM-1 in major group HRVs and of PVR in PVs. However,

the proposed mechanism by which canyon-binding receptors initiate the

instant uncoating of the virus (Rossmann et al., 2002) may not to be valid for

EV1. Instead, the integrin-interaction may inhibit the instant disassembly of

EV1 particles to ensure that the release of viral genome takes place after

internalization of the virus into the cells.

The significant differences between EV1 and collagen in their binding to

α2β1 integrin strengthen the view that viruses have, during evolution,

maximised their capability to compete with physiological ligands. To ensure

the efficient attachment to the cell surface, EV1 has taken benefit of the

higher affinity for the receptor compared to the natural ligand and of the

ability of the virus to bind several integrin molecules simultaneously.

Moreover, EV1 has exploited the signalling properties of the integrins to

facilitate its internalization and replication. On the other hand, the signalling

triggered by the receptor after interactions with the virus can also prepare the

host cell for the viral invasion (Pelkmans and Helenius, 2003).

To cross the membrane barrier as well as to evade the host immune system, it

is advantageous for viruses to utilize the endocytic machinery. Unlike many

other picornaviruses, EV1 does not utilize clathrin-mediated uptake. Instead,

Conclusions

75

it can be internalised into caveosomes through cell surface caveolae and/or

through another, still undefined, internalization route that may involve non-

caveolar lipid rafts. Determination of particular cellular components and

molecular mechanisms involved in EV1 uptake into caveosomes will be of

great importance in the future studies.

EV1 may remain in caveosomes until the uncoating and replication of the

virus take place. Thus, it differs from other known caveosomal ligands, which

traffic from caveosomes to the ER or to the Golgi (Pelkmans et al., 2001; Le

and Nabi, 2003). Therefore, it will be relevant to investigate the specific

mechanisms of viral uncoating and viral RNA release in/from caveosomes. In

addition, EV1 provides an excellent instrument for solving the missing links

of the caveosomal pathway because the viral genome is easy to manipulate

and, as shown here, the direct fluorescent labelling of EV1 capsid proteins

provides a useful tool to follow the dynamic process of virus uptake into live

cells. Real-time live microscopy of fluorescently labelled virus particles could

also be applied to the elucidation of currently unknown endocytic routes of

other picornaviruses.

EV1, like all viruses, survives during the interaction with host cells by

controlling the cell’s function by multiple mechanisms, including the

regulation of host cell protein translation and gene expression. cDNA arrays

are advantageous for identifying universal patterns in cellular gene expression

during virus infections. In the future, cDNA arrays may also have a

considerable impact on the diagnosis and therapy of infectious diseases

(Manger and Relman, 2000). In spite of significant benefits of cDNA array

techniques, additional investigations are required to resolve which of those

host genes shown to be regulated during EV1 infection are involved in EV1-

host cell interactions, in general mechanisms of viral pathogenesis and in host

cell defence.

Viruses have evolved to utilize the existing cellular molecules and functions

for infection. Therefore, the studies on virus infections have provided

solutions to several phenomena in molecular and cell biology. The findings

reported here demonstrate that EV1 can serve as such a "model" virus to

Conclusions

76

illuminate virus - host cell interactions. Hopefully, the findings presented in

this thesis will promote studies of virus-receptor interactions in general as

well as more detailed investigations of currently poorly known cellular

pathways.

Acknowledgements

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ACKNOWLEDGEMENTS

The studies of this thesis were carried out at the Department of Virology, HaartmanInstitute, University of Helsinki. I would like to thank Professor Emeritus AnttiVaheri, who was the Head of the Department of Virology at the time of this study, aswell as the present Head of the Department, Professor Kalle Saksela, for providing acreative and enthusiastic environment for research. Professor Carl Gahmberg, theHead of former Department of Biochemistry, where I started my studies inbiochemistry in 1994, is warmly thanked for leading me to the fascinating world ofbioscience.

I want to express my gratitude to the supervisor of the thesis, Professor Timo Hyypiä,who has encouraged me to use my imagination for solving the scientific problems,taught me an open-minded attitude towards new ideas. After setbacks, he has alwayswisely suggested a new approach.

The reviewers of my thesis, Professor Elina Ikonen and Docent Maarit Suomalainen,are thanked for fruitful comments on the manuscript that greatly helped to improve it.Professors Jyrki Heino and Antti Vaheri are thanked for giving valuable ideas andcomments on my studies when acting as members of my thesis committee. I also wishto thank Professor Jyrki Heino for collaboration and for giving me an opportunity tovisit several times his laboratory at the University of Jyväskylä.

I thank our collaborator Docent Varpu Marjomäki at the University of Jyväskylä forher positive attitude and for stimulating discussions. Also, I would like to thank Dr.Paula Upla, who helped me to carry out some important experiments for the last paperof the thesis during my maternity leave. I wish to express my gratitude to ProfessorAri Helenius for kindly letting me to work in his laboratory at the Swiss FederalInstitute of Biotechnology Zürich for two months, and for giving so much of hisvaluable time for supervising me in scientific issues and in writing of ourcollaborative article. Dr. Lucas Pelkmans, who worked in the laboratory during myvisit, is thanked for showing me the secrets of real-time live microscopy. ProfessorHolland Cheng and Dr. Li Xing from Karolinska Institute, Sweden, as well asProfessor Mark Johnsson and Dr. Mikko Huhtala at the Åbo Akademi are greatlythanked for collaboration in cryo-EM studies of EV1-α2I domain. I would like tothank Holland especially for inspiriting discussions about virus uncoating. Li andMikko, thank you for patience to answer to my never-ending questions on structuralbiology and for your company in several meetings! I also want to thank othercollaborators including Pasi Huttunen, Johanna Ivaska, Jarmo Käpylä, HeliMatilainen, Liisa Nissinen, Hilkka Reunanen, and Kirsi Vuorinen. In addition, I haveenjoyed discussions with experienced picornavirologists, including Professor TapaniHovi, Docent Merja Roivainen and Professor Glyn Stanway.

Acknowledgements

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I would like to thank for all the former and present members of our group, includingAnnu, Åse, Camilla, Olya, Hannamari, Maaria, Pia, Veera, Heli, Päivi, Tommi, Anne,Juhana, Pasi, Timo P, Tuija, Tiiu, Seija, Leena K, Leena P, Maarit, Henrietta, Heidi....and many others. Picorna-girls, I have enjoyed your hilarious company in thelaboratory and elsewhere! Heli, thank you for positive friendship. In addition, Maariaand Tiiu, thank you for helping me in my work.Leena Kostamovaara is specially thanked for helpful attitude during these years. LauriYlimaa is thanked for helping me with processing figures for the original publicationsand for this thesis. I wish to thank Professor Malcolm Richardson for revising thelanguage of my thesis. Finally, I want to thank all people at the Department ofVirology for a cosy working atmosphere.

My friends, relatives and family, you have given balance to my life outside thelaboratory. Special thanks to good old friends from the childhood, especially to Mari,to friends from the Department of Biochemistry, and to Marikka and Janne. I wouldlike to thank Titta for being such a marvellous friend, also through "delightful anddreadful days" in science! My wonderful parents-in-law, Ukko ja Ulla, thank you forhelping me to organize time to complete my thesis.

My parents, Äiti ja Isi, thank you for always being there for me. Thank you forsupporting me and believing in me in everything I have decided to do.

My dearest thanks to Tatu and Tuuve. Tatu, you have encouraged me with patience tocomplete this thesis and to overcome all the scientific troubles. Thank you for yourlove and for sharing the happiest moments of my every-day life, especially the mostjoyful time together with our little "humorist" Tuuve.

This thesis was financially supported by the Academy of Finland, the HelsinkiGraduate School of Biotechnology and Molecular Biology, the Finnish CulturalFoundation, Kymenlaakson rahasto of the Finnish Cultural Foundation, the PauloFoundation, Virustautien tutkimussäätiö, the European Molecular BiologyOrganization, the Biomedicum Foundation and the Sigrid Juselius Foundation.

Helsinki, May 2005

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