Advances in Virus Research, Volume 77

ADVISORY BOARD DAVID BALTIMORE ROBERT M. CHANOCK PETER C. DOHERTY H. J. GROSS B. D. HARRISON BERNARD MOSS ERLING NORRBY J. J. SKEHEL M. H. V. VAN REGENMORTEL

Academic Press is an imprint of Elsevier 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 32 Jamestown Road, London, NW1 7BY, UK Linacre House, Jordan Hill, Oxford OX2 8DP, UK First edition 2010 Copyright # 2010, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (þ44) (0) 1865 843830, fax: (þ44) (0) 1865 853333; e-mail: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://www.elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-12-385034-8 ISSN: 0065-3527 For information on all Academic Press publications visit our website at elsevierdirect.com Printed and bound in USA 10 11 12 10 9 8 7 6 5 4 3 2 1

CHAPTER

1 Recent Developments in Understanding Dengue Virus Replication Silvio Urcuqui-Inchima,* Claudia Patin˜o,* Silvia Torres,* Anne-Lise Haenni,*,† and Francisco Javier Dı´az*

Contents

I. Introduction II. Dengue Disease A. The agent B. Eco-epidemiology C. Clinical manifestations D. Pathogenesis III. DENV Genome and DENV Proteins IV. Cell Cycle of DENV A. Entry and fusion complex formation: E protein and cell receptors B. Virus adsorption and fusion to the endosome membrane C. Intracellular transport of the viral genome D. Genome expression E. Genome replication F. Maturation and release of DENV from the host cell V. siRNAs and New Strategies to Control DENV Replication

3 4 4 5 5 6 8 11 11 19 19 20 21 24 26

* Grupo de Inmunoviologı´a, Sede de Investigacio´n Universitaria, Universidad de Antioquia, Medellı´n, {

Colombia Institut Jacques Monod, Paris, France

Advances in Virus Research, Volume 77 ISSN 0065-3527, DOI: 10.1016/S0065-3527(10)77001-9

#

2010 Elsevier Inc. All rights reserved.

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VI. Conclusions Acknowledgments References

Abstract

27 28 28

Dengue is the most important cause of mosquito-borne virus diseases in tropical and subtropical regions in the world. Severe clinical outcomes such as dengue hemorrhagic fever and dengue shock syndrome are potentially fatal. The epidemiology of dengue has undergone profound changes in recent years, due to several factors such as expansion of the geographical distribution of the insect vector, increase in traveling, and demographic pressure. As a consequence, the incidence of dengue has increased dramatically. Since mosquito control has not been successful and since no vaccine or antiviral treatment is available, new approaches to this problem are needed. Consequently, an in-depth understanding of the molecular and cellular biology of the virus should be helpful to design efficient strategies for the control of dengue. Here, we review the recently acquired knowledge on the molecular and cell biology of the dengue virus life cycle based on newly developed molecular biology technologies.

ABBREVIATIONS aa ADE C Cdc42 cHP CS DAR DC DC-SING DENV DF DHF ds DSS E eEF eIF ER FcgR

amino acid antibody-dependent enhancement capsid/coat protein cell division cycle 42 capsid hairpin cyclization sequence downstream of the AUG region dendritic cells dendritic cell-specific intercellular adhesion molecule 3grabbing non-integrin dengue virus dengue fever dengue hemorrhagic fever double-stranded dengue shock syndrome envelope eukaryotic elongation factor eukaryotic initiation factor endoplasmic reticulum receptor of the Fc portion of IgG

Dengue Virus Replication

GFP HLA HMEC HSP IF IFN IL LC3-II LD M MT MTase NC NS nt NTPase ORF PABP prM Rab 5 Rac1 RC RER RdRp RTPase siRNA SL sgRNA ss TGN TNF UAR UTR V-ATPase WHO

3

green fluorescent protein histocompatibility-linked antigen human microvascular endothelial cell heat-shock protein intermediate filament interferon interleukin light chain 3 form II of the microtubule-associated protein 1 lipid droplet matrix protein microtubule methyltransferase nucleocapsid nonstructural nucleotide nucleoside triphosphatase open reading frame poly(A)-binding protein precursor of M Rabatin-5 ras-related c3 botulinum toxin substrate 1 replication complex rough ER RNA-dependent RNA polymerase RNA 50 -triphosphatase small interfering RNA stem-loop subgenomic RNA single-stranded trans-Golgi network tumor necrosis factor upstream AUG region untranslated region vacuolar-ATPase World Health Organization

I. INTRODUCTION Although dengue only became a major global medical concern within the last few decades, disease that might have been caused by dengue was reported in China as early as the seventh century. Since then, epidemics that may have been caused by dengue have sporadically appeared with

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outbreaks in the seventeenth century in Central America and in Philadelphia in the eighteenth century. In the twentieth century, epidemics were reported in Southeast Asia and the Pacific, favored by increased expansion of the dengue virus (DENV) mosquito vector, Aedes aegypti. This was accompanied by circulation of the four DENV serotypes (known as hyperendemicity) and the appearance of dengue hemorrhagic fever (DHF). By the 1970s, DHF had become a major cause of death among children in regions where DHF occurred. Since then, epidemics have become more frequent and more intense, and cover an ever increasing geographical area. Although before the 1980s Africa had not experienced major epidemics, it now harbors all four DENV serotypes (reviewed in Gubler, 1998). DENV continues to be a public health problem in tropical and subtropical countries. With the recent increase in dengue epidemics, interest in attempts to control dengue has expanded. It has become urgent to control dengue, because to date, neither animal model, vaccine, nor therapeutic measures are available to counteract DENV and the disease. One of the control measures adopted had been vector control. However, although this strategy seemed to be highly promising, it was later neglected, resulting in renewed epidemics in various parts of the world. Safe and efficient strategies will need to be developed to control the vector and will require the active participation of local communities. The aim of the present review is to bring together recent knowledge acquired on the molecular and cell biology of the life cycle of DENV in the hope that it might lead to a better understanding of dengue diseases. References to recent review articles related to the field of research presented here can be found throughout the present review.

II. DENGUE DISEASE A. The agent Dengue is the most widespread arthropod-borne viral disease of humans and is caused by DENV. DENV is a spherical, enveloped virus with a monopartite singlestranded (ss) positive-sense RNA [ssRNA(þ)] of the family Flaviviridae, genus Flavivirus (reviewed in Mukhopadhyay et al., 2005). Four DENV serotypes are recognized, DENV-1 to DENV-4 that possess 67–73% identity at the nucleotide (nt) level and 69–78% identity at the amino acid (aa) level. Three to six subtypes (genotypes) can be distinguished within each serotype (Diaz et al., 2006). Monoclonal antibodies made it possible to identify epitopes that are serotype-specific, DENV complex-specific, and flavivirus group-reactive epitopes as well as intermediate categories.

Dengue Virus Replication

5

All serotypes have the same transmission cycles and cause similar clinical manifestations, although serotype- and strain-related differences in virulence exist. Infection by one serotype is believed to confer life-long immunity to only that serotype (reviewed in Rico-Hesse, 2010).

B. Eco-epidemiology DENV is vectored by Aedes mosquitoes. Humans and A. aegypti are generally the only vertebrate reservoir and vector respectively, but forest and rural cycles involving nonhuman primates and Aedes species other than A. aegypti exist. A. aegypti breeds mainly in human-made containers and about 2.5 billion people are at risk in regions where A. aegypti is endemic (www.who.int/mediacentre/factsheets/fs117/en/). A general tendency toward increased frequency and magnitude of dengue outbreaks has occurred recently. The reasons invoked are demographic, cultural, environmental, and political (Guha-Sapir and Schimmer, 2005; Wilder-Smith and Gubler, 2008). Prominent factors implicated are increase in global population and in air travel, and rapid and unplanned urbanization. This has resulted in the simultaneous circulation of multiple DENV serotypes, known as hyperendemicity. Other factors include the increasing use of disposable products that become mosquito breeding sites, decay in public health infrastructure, insecticide resistance, and global warming.

C. Clinical manifestations Infection by DENV results in clinical outcomes ranging from asymptomatic to fatal. Traditionally, dengue cases have been classified as undifferentiated fever, dengue fever (DF), and DHF; DHF has been further divided in cases with or without shock, the first being designated as dengue shock syndrome (DSS). A more recent classification divides the disease into dengue (with or without warning signs) and severe dengue; the second category includes cases with significant plasma leakage, hemorrhage, or organ impairment, comprising the old categories of DHF and DSS, as well as other severe forms of DENV-induced disease that affect the brain, liver, heart, or other organs [World Health Organization (WHO), 2009]. Although this latter classification promises improved reporting of cases and better medical management, here we will often use the more familiar DF, DHF, and DSS designations. At least half of the DENV infections are asymptomatic or mild. Yet, DF is an acute disease characterized by fever and pains in the head, eyes, muscles, and joints. Lymphadenopathy, rash, nausea, and minor hemorrhages also occur. The course is usually benign and self-limited. A variable proportion of cases evolves into DHF, a condition defined by fever,

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thrombocytopenia (< 100,000 platelets/mm3), hemorrhages (or a positive tourniquet test) and increased vascular permeability. DHF usually appears three to eight days after fever begins. It is often preceded by acute abdominal pain and is commonly characterized by hepatomegaly and bleeding from the skin, mucosae, and the gastrointestinal tract. Patients with most severe DHF develop hypotension and other signs of shock that characterize DSS, and these can lead to death unless prompt fluid replacement therapy is administered to them (reviewed in Gubler, 1998; Guzman and Kouri, 2002).

D. Pathogenesis No appropriate animal models exist to study DENV infections. Mice can be infected intravenously but neither viremia nor disease develops. Monkeys develop an infection but viremias are low and no disease is observed. Thus, human infections have supplied most of the information available. Human infection begins in the skin with the injection of DENV by the bite of an infected mosquito. DENV probably initially replicates in dendritic cells (DC) or macrophages (Marovich et al., 2001; Wu et al., 2000) and then migrates to regional lymph nodes from where it spreads to other lymphoid organs through the bloodstream. It is detected in the plasma and in peripheral blood mononuclear cells. Monocytes, macrophages, and DCs are major targets of DENV replication, although Kupffer cells, hepatocytes, and lymphocytes have also been implicated as target cells (Halstead et al., 1977; King et al., 1999). Antibody response to DENV infection is different in the first (primary) and subsequent (secondary) infections. In the former, a strong IgM response appears by the end of the febrile period and lasts two to three months; IgG appears shortly after IgM, increases moderately, and is maintained for years. These antibodies are predominant against the infecting serotype, but a low titer crossreactive (heterotypic) response is also observed. After a subsequent infection, IgG levels increase more rapidly and to higher levels than in primary infections; IgM response is weaker and declines rapidly; antibodies are mostly heterotypic and titers are usually higher against the first infecting serotype (Vaughn et al., 1997). Antibodies recognize both structural and nonstructural DENV proteins, but only those reactive with the E (envelope), M (matrix) and nonstructural (NS) 1 proteins have neutralizing properties (reviewed in Pierson and Diamond, 2008). The duration of protection against re-infection with DENV remains unclear. Humans challenged with the homologous strain were completely immune to re-infection for as long as 18 months. Temporal cross-protection to challenge with a heterologous serotype was demonstrated two to nine months after the primary infection.

Dengue Virus Replication

7

DENV also elicits specific CD4þ and CD8þ T cell immune responses. After primary infection, most circulating T lymphocytes are serotypespecific, but following a second infection with a different serotype, cross-reactive memory T cells predominate and more frequently recognize highly conserved proteins, especially NS3. Both CD4þ and CD8þ T lymphocytes can lyse DENV-infected cells. DENV-specific lymphocytes release mostly Th1 cytokines, which could be important factors in viral clearance as well as in the pathogenesis of DHF (reviewed in Mathew and Rothman, 2008). Features of DHF/DSS are increased vascular permeability, bleeding, and hepatic compromise. Fatal cases present bleeding in the viscera and mucosal membranes, liquid collection in serous cavities, and midzonal necrosis in the liver. Although mostly a vascular illness, vascular endothelial damage is not observed in DHF, suggesting a functional rather than anatomical alteration in the endothelium (reviewed in Gubler, 1998; Guzman and Kouri, 2002; Trung and Wills, 2010). Severe forms of DHF/DSS are associated with young age, female gender, non-African ancestry, and preexistence of immunity against other serotypes. Most severe cases are associated with a heterotypic antibody response confirming secondary infection as the main risk factor for DHF/DSS (Burke et al., 1988; Kliks et al., 1989; Sangkawibha et al., 1984). The finding of preexisting heterotypic antibodies in cases of DHF is linked to a phenomenon designated as antibody-dependent enhancement (ADE). It consists of increased replication of certain viruses in cell culture by the addition of immune sera at dilutions beyond the neutralization endpoint. This led to the ‘‘immune enhancement hypothesis,’’ whereby nonneutralizing antibodies remaining after the first dengue episode bind to the new infecting serotype enhancing virion penetration into monocytes and macrophages by interacting with the receptor for the Fc portion of IgG, known as FcgR (see Section IV.A). This phenomenon would lead to enhanced viral replication and release of soluble factors that would mediate increased vascular permeability and hemostatic disorder (Halstead and O’Rourke, 1977; reviewed in Halstead, 1981). In spite of considerable efforts, the mechanism involved in ADE remains unclear. Dejnirattisai et al. (2010), using monoclonal antibodies against the precursor of M (prM), showed that these antibodies, even though they crossreact with the various DENV serotypes, are unable to block DENV infection, but on the contrary, promote ADE. This interaction is likely due to some uncleaved prM remaining on the virus surface of extracellular virions that would allow their recognition by the monoclonal antibody. Cellular immune factors are also involved: DENV-specific CD4þ cells release cytokines such as interferon (IFN)-g, interleukin (IL)-2, tumor necrosis factor (TNF)-a, and TNF-b (Gagnon et al., 2002). TNF-a and IL-2 can induce plasma leakage, and IFN-g enhances TNF-a production

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by monocytes. IFN-g also upregulates the expression of class II histocompatibility-linked antigens (HLAs) and of the receptor FcgR (the Fc portion of IgG) in monocytes; this could enhance DENV antigen presentation and antibody-mediated uptake of the virus, respectively. Increased levels of TNF-a, IL-6, IL-8, IL-10, and of soluble IL-2 and TNF receptors occur in DHF (Chang and Shaio, 1994; Hober et al., 1993). Immunity against a different serotype is neither necessary nor sufficient for DHF/DSS development. Severe forms of DENV can occur in primary infections (Barnes and Rosen, 1974; Morens et al., 1987), whereas in other settings, DHF/DSS was never observed in spite of frequent sequential infections by different serotypes. Viral risk factors associated with severe forms of DENV-induced disease include the infecting serotype, subtype, and strain, the time lapse between infections and the order of the serotypes in successive infections. A compelling demonstration of the importance of the virus strain in the severity of dengue came from the Americas where epidemic DHF emerged abruptly in 1981 when a Southeast Asian subtype of DENV-2 was introduced. This subtype then spread across the Caribbean and Central and South America, displacing the prevalent ‘‘American subtype’’ of DENV-2, which had been associated with mild DF cases only (Gubler and Clark, 1995; Watts et al., 1999). The time of arrival of the Southeast Asian subtype coincided with the emergence of DHF/DSS in each of these regions. At the phenotypic level, certain DENV isolates are more infectious and are disseminated more efficiently by A. aegypti mosquitoes than other isolates (Gubler et al., 1978). At the genotypic level, consistent differences between isolates of two DENV-2 subtypes were detected in several viral genes involved in severity (Cologna and RicoHesse, 2003). Thus, even minor differences at the molecular level can affect the clinical outcome. Yet, the specific mechanisms involved are unresolved, because the role of most viral proteins remains elusive. Elucidation of these roles is of utmost importance to understand the molecular basis of virulence and attenuation, which is fundamental to the design of the longawaited safe and efficacious vaccines for dengue. This review will now focus on the molecular aspects of DENV replication to describe the viral and cell factors that could be modulating the clinical and epidemiological behavior of dengue.

III. DENV GENOME AND DENV PROTEINS The ssRNA(þ) of DENV (Fig. 1B) acts directly as mRNA for the synthesis of the viral polyprotein (Fig. 1C). The genome is approximately 11 kb, bears a type I cap structure (m7GpppAmG) at its 50 end, but lacks a 30 poly

9

Dengue Virus Replication

A Top loop

SLA Side stem loop

cHP

U bulge

*

5⬘–3⬘UAR

5⬘–3⬘CS

5⬘ 3⬘

B

3⬘SL

Replication

Top loop

Side stem loop ORF

U bulge

SLA

~10.8 kb 5⬘UAR

3⬘UAR 5⬘CS

* SLB

3⬘CS

cHP

C

3⬘SL 3⬘UTR

Translation

5⬘UTR

NS2B/NS3 Signal peptidase Furin

~3400 aa

Unknown Polyprotein

NH3 C

PrM

E

NS1

NS2A NS2B

2K MTase

Hel

Prot NS3

NS4A

NS4B

RdRP

COOH

NS5

FIGURE 1 The DENV genome: (A) schematic diagram of DENV genome circularization; (B) organization of the 50 and 30 UTRs; (C) polyprotein. (A, B) The viral ssRNA (þ) genome is  11 kb long. The 50 UTR contains the large stem-loop A (SLA), the promoter for NS5, the RNA-dependent RNA polymerase (RdRP), followed by stem-loop B (SLB) that contains the 50 upstream AUG region (50 UAR) and the translation initiation codon (*). The 30 UAR is complementary to the 50 UAR. Another stem-loop hairpin structure (cHP) located within the C protein-coding region enhances selection of the 50 initiation codon. The 30 UTR contains conserved sequences such as the 30 stem-loop (30 SL) that includes the 30 UAR. The 30 cyclization sequence (30 CS) lies upstream of the 30 SL; it is complementary to the 50 CS present in the gene encoding the C protein. The predicted 50 –30 UAR and 50 –30 CS sequences are in gray, and hybridization between these regions is necessary for genome cyclization and RNA synthesis. (C) The viral genome possesses one ORF coding for a polyprotein. The genes for the structural proteins are the capsid (C), the precursor of membrane protein (PrM) and E, followed by the genes for the NS proteins (NS1, NS2A, NS2B, NS3, NS4A, NS4B, and NS5). The polyprotein is processed in the cytoplasm by NS2B–NS3 (closed circles), by a signal peptidase (small arrow), or by an as yet unknown proteinase (open triangle) in the lumen of the ER, and PrM is processed by furin (closed triangle) in the cellular secretory pathway.

(A) tail. It encodes a single open reading frame (ORF) flanked by highly structured 50 and 30 untranslated regions (UTRs) of about 100 and 400 nts, respectively (Fig. 1B); these regions are important for translation regulation, viral RNA replication (Fig. 1A), and severity of infection

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Silvio Urcuqui-Inchima et al.

(Leitmeyer et al., 1999; Edgil et al., 2003; Holden and Harris, 2004; reviewed in Markoff, 2003; Bartenschlager and Miller, 2008). The cap structure is added cotranscriptionally by the virus-encoded NS5 protein which contains in a single domain both (guanine-N7)- and (adenosine-20 -O)-methyltransferase (MTase) activities, and an as yet unidentified guanylyltransferase domain (Egloff et al., 2007; reviewed in Davidson, 2009). The RNA 50 -triphosphatase (GMP-k) domain of NS3 is also involved in capping the viral RNA; although the precise role of NS3 in this process has not been well characterized, several studies have shown that the NS3 RTPase and the NS5 MTase work together removing the terminal g-phosphate and performing sequential guanine-N7 and adenosine-20 -O methylation, respectively (Egloff et al., 2002; Wengler and Wengler, 1993). The cap structure presumably stabilizes the mRNA and allows efficient translation (reviewed in Furuichi and Shatkin, 2000). The ORF is translated as a single polyprotein of 3387–3392 aa that undergoes co- and posttranslational cleavages by viral and host cell proteinases (Fig. 1C). In the region of the ORF encoding the capsid (C) protein, a stemloop hairpin structure designated as capsid hairpin (cHP) involved in codon start selection, and viral RNA replication has been identified (Clyde and Harris, 2006; Hahn et al., 1987), followed by a 50 cyclization sequence (CS) complementary to a 30 CS in the 30 UTR (Fig. 1A and B; Hahn et al., 1987). The ORF of over 10 kb comprises from its 50 to 30 end, information for the synthesis of three structural proteins C, prM, and E and the seven NS proteins (NS1-NS2A-NS2B-NS3-NS4A-NS4B-NS5; Fig. 1C), a feature common to all members of the genus Flavivirus. Yet, size heterogeneity exists between the noncoding regions, particularly in the 30 UTR (Bryant et al., 2005; Medeiros et al., 2007; Wallner et al., 1995). The 50 UTR (Fig. 1A and B) bears the stem-loop (SL) A, and the SLB that ends with the initiator AUG codon (Brinton and Dispoto, 1988). The SLA acts as promoter element recognized by NS5, which contains an RNAdependent RNA polymerase (RdRp) domain for RNA synthesis (Filomatori et al., 2006). It harbors a side stem-loop and a top loop, in addition to a U bulge (Lodeiro et al., 2009) that are critical for DENV RNA replication. SLB contains upstream of the initiator AUG codon, a sequence known as 50 UAR (upstream AUG region), that is complementary to a sequence located at the 30 end of the viral genome (Alvarez et al., 2005a; Brinton and Dispoto, 1988). The 30 UTR (Fig. 1A and B) includes three regions, a variable region immediately after the termination codon of the ORF, followed by a core region, and finally a 30 -terminal region. The latter includes a conserved 30 SL of  96 nt (Brinton et al., 1986; Grange et al., 1985; Mohan and Padmanabhan, 1991). The 30 SL contains a conserved 30 UAR sequence that is complementary to the 50 UAR located in the 50 UTR (Fig. 1A;

Dengue Virus Replication

11

Alvarez et al., 2005a); a distal flavivirus-conserved CACAG Box has also been described (Chen et al., 1997; Filomatori et al., 2006; reviewed in Markoff, 2003). The genome contains upstream of the 30 UAR, a conserved 30 CS (Hahn et al., 1987) complementary to the conserved 50 CS and which together with the 50 UAR and 30 UAR is involved in RNA cyclization (Clyde and Harris, 2006; Hahn et al., 1987; Men et al., 1996). Moreover, a sequence located downstream of the initiation codon, designated as 50 downstream AUG region (DAR), is involved in DENV replication (Friebe and Harris, 2010). Co- and posttranslational cleavage of the DENV polyprotein is achieved by the viral-coded serine proteinase NS2B–NS3, by the cell convertase furin, a proteinase of the trans-Golgi network (TGN), and by a cell signal peptidase (Falgout et al., 1991; Cahour et al., 1992; reviewed in Lindenbach and Rice, 2003). Polyprotein cleavage sites, conserved and unique motifs, important cysteine residues, and potential glycosylation sites have been characterized (reviewed in Gubler et al., 2007; Mukhopadhyay et al., 2005). Table 1 provides a summary of the major activities so far identified for the viral proteins.

IV. CELL CYCLE OF DENV A. Entry and fusion complex formation: E protein and cell receptors Monocytes, macrophages, B and T lymphocytes, hepatocytes, endothelial cells, epithelial cells, DC, and fibroblasts are all potential targets for DENV infection and replication, and viral antigens have been detected in liver, spleen, lymph node, thymus, kidney, lung, and skin cells (Upanan et al., 2008). Hence, the virus can replicate in a wide spectrum of cells, which might explain its ability to enter via various receptors, as discussed later. Receptor binding and membrane fusion characteristics of DENV entry have been studied in living cells by real-time fluorescence microscopy and by direct biochemical or genetic analyses. Yet, the reports published on virus entry and on the molecules involved in this process, are controversial and may vary with the cell model used. Considerable evidence suggests that the first step in a primary infection is attachment of the viral glycoprotein E which is present on the surface of mature virions as a homodimer, to cell receptors on the surface of permissive cells (Fig. 2, step 1). The E protein consists of three domains. Domain I is in the N-terminal region and provides an organizational structure. Domain II or central domain is involved in dimerization and contains a hydrophobic fusion loop essential for fusion of E to the cell membrane. Domain III

TABLE 1

Characteristics and functions of the DENV proteins

Viral Size protein (kDa) Functions

C

11

– N- and C-termini have charged residues – Central hydrophobic region that associates with membrane – C-terminal hydrophobic sequence that acts as a signal peptide for translocation to ER – Has nuclear localization signal and is found in the cytoplasm and nucleus; its role in the nucleus is unknown – Forms NCs with viral RNA; its dimerization triggered by viral RNA is required for virus assembly – Interacts with the apoptotic protein DAXX inducing apoptosis – Interacts with human Sec3, a member of the exocyst complex, a transcription and translation repressor of flavivirus that can retard infection

prM

34

References

Wang et al. (2002), Ma et al. (2004) Ma et al. (2004) Markoff et al. (1997) Tadano et al. (1989), Wang et al. (2002), Bulich and Aaskov (1992), Tsuda et al. (2006), Sangiambut et al. (2008) Ma et al. (2004), Ku¨mmerer and Rice (2002), Kiermayr et al. (2004), Lopez et al. (2009), reviewed in Bartenschlager and Miller (2008) Netsawang et al. (2010) Bhuvanakantham et al. (2010)

– Precursor of membrane protein M, cleaved into Elshuber et al. (2003) pr þ M (26 þ 8 kDa) by furin located in TGN; required for infectivity – Liberated from polyprotein by host signalase located in reviewed in Lindenbach and Rice (2003) the ER – Importance of His39 in the morphology of virus, and Pryor et al. (2004) secretion and entry of virus into cell

– Interaction with E forms spikes on the virus surface, and is required for proper folding and secretion of E – Interaction with V-ATPase required for proper egress of virus – pr leaves virions after secretion of particles and exposure to neutral pH – By interacting with E, the pr peptide prevents fusion of E to cell membranes – pr blocks membrane fusion by binding to virus at acidic pH – Binds to the claudin-1 tight junction membrane protein that protects cells from the environment; interaction is required for virus entry E

50

– Intracellularly linked to prM forming heterodimers that protect E from premature acidification during transit through Golgi – Forms outer glycoprotein shell of virus – Major target for neutralizing antibodies – Cleavage of prM produces reversible conformational change in E – Protected by pr retained on particle from premature fusion to cell – Interaction with chaperones (Bip, calnexin, calreticulin) required for virus production

Duan et al. (2008), Li et al. (2008), reviewed in Lindenbach and Rice (2003) Duan et al. (2008) Li et al. (2008), Yu et al. (2008a) Zhang et al. (2003), Li et al. (2008), Yu et al. (2008a, 2009) Yu et al. (2009) Gao et al. (2010)

Guirakhoo et al. (1993), Konishi and Mason (1993), reviewed in Perara et al. (2008) reviewed in Lindenbach and Rice (1997, 2003) reviewed in Halstead (1988), Green and Rothman (2006) Li et al. (2008), Yu et al. (2008a) Li et al. (2008), Yu et al. (2008a) Limjindaporn et al. (2009) (continued)

TABLE 1

(continued)

Viral Size protein (kDa) Functions

– Specific amino acids determine hemorrhagic disease or encephalitis – Low pH in Golgi induces change in E structure important for formation of fusion complexes – Interacts with vacuolar-ATPase important for entry and egress of virus at low pH – E glycosylation decreases infectivity and increases virus release – Natural killer (NK) cell-activating receptor NKp44 interacts with E, leading to activation of NK cells and destruction of virally infected cells – Interaction with chaperones in ER facilitates folding and assembly of DENV proteins – Its transmembrane domain is responsible for retention and assembly of E on ER NS1

46

– In flaviviruses, proposed to act early in replication cycle – May define pathogenesis – Possibly interacts with NS4A, thereby participates in viral RNA replication

References

Barker et al. (2009) reviewed in Heinz and Allison (2003) Duan et al. (2008) Lee et al. (2010), Duan et al. (2008) Hershkovitz et al. (2009) Limjindaporn et al. (2009)

Hsieh et al. (2010)

Lindenbach and Rice (1997, 1999), Muylaert et al. (1997) Falconar (1997) Lindenbach and Rice (1999)

NS2A 22

– With NS4A and NS4B, is involved in IFN resistance and blocks IFN-b signal

NS2B 14

– Cofactor of NS3, forming serine proteinase activity in Westaway et al. (2010) trans; acts in cis to cleave NS2B–NS3 Falgout et al. (1991) – Required for proteolytic processing of DENV nonstructural proteins

NS3

– Has N-terminal region with serine-proteinase acivity and C-terminal region with NTPase, RNA helicase, and 5’ RNA triphosphatase activities – Cleavage activity retained in NS2B–NS3 heterodimer – NS5 stimulates NS3 activities – Required for proteolytic processing of DENV nonstructural proteins – Binds to La protein

70

Munoz-Jordan et al. (2003)

Wengler and Wengler (1993), Li et al. (1999), Sampath et al. (2006), Luo et al. (2008a,b) Leung et al. (2001), Bera et al. (2007) Yon et al. (2005) Falgout et al. (1991) Garcia-Montalvo et al. (2004)

NS4A 16

– C-terminal region known as 2K fragment; involved in Zou et al. (2009), Miller et al. (2007) regulating membrane rearrangements – Associates with membranes through its hydrophobic reviewed in Lindenbach and Rice (2003) regions – Probably involved in RNA replication Miller et al. (2007) – Induces membrane rearrangements resembling virus-induced structures

NS4B 27

– Blocks IFN-induced signal-transduction cascade – Interacts with NS3 and dissociates it from ssRNA

Munoz-Jordan et al. (2003) Umareddy et al. (2006) (continued)

TABLE 1

(continued)

Viral Size protein (kDa) Functions

– Downregulates expression of STAT2 – Participates in formation of RCs NS5

103

References

Jones et al. (2005) Miller et al. (2006)

– Has MTase (N-terminus) and RdRp (C-terminus) motifs

Ackermann and Padmanabhan (2001), Egloff et al. (2002), Yap et al. (2007); Selisko et al. (2010) – Possesses two nuclear localization signals for transport Brooks et al. (2002)

to nucleus – Is a nuclear phosphoprotein; only hypophosporylated NS5 is located in cytoplasm where it interacts with NS3 – Modulates enzymatic activities of NS3 – Possesses a nuclear export signal and interacts with CRM1 for export to cytoplasm – Participates in methylation of viral RNA cap structure – Binds to La protein – Its interaction with STAT2 required for IFN signaling, reduces level of expression of STAT2. Inhibits IFN-a signaling by binding to the STAT2 and inhibits its phosphorylation

Kapoor et al. (1995) Rawlinson et al. (2009) Yon et al. (2005) Rawlinson et al. (2009) Egloff et al. (2002), Geiss et al. (2009) Garcia-Montalvo et al. (2004) Ashour et al. (2009), Mazzon et al. (2009)

1.

Cell protiens

2.

Viral protiens

PABP

PrM

elF4G

E

elF4E

NS2A

Furin

NS3

Receptor

3.

NS4B

Ribosome

NS5

TGN

4. NC 5. RNA(+)

10.

11.

6. RNA(−) 9. 8.

N

G

ER 7.

FIGURE 2 Schematic representation of the DENV life cycle. The DENV E protein is the major component of the virion surface. (1) Attachment: The initial step in the viral life cycle is attachment of the E protein to a cellular receptor (forming a fusion complex) such as heparan sulphate, a mannose-specific C-type lectin, but a specific receptor for internalization of the virus into host cells has not been identified. (2) Endocytosis: Following receptor binding, the virus is internalized into the cells through clathrinmediated endocytosis transporting the virus particles to endosomes (3) Fusion of membranes: In the cytoplasm, acidification of the endosome lumen induces structural changes in E and promotes fusion between the virus particle and the endosomal membrane. (4) Uncoating: A fusion pore is formed, the nucleocapsid (NC) is delivered into the cytoplasm, and after uncoating, the viral RNA (red) is released from the NC into the cytoplasm. (5) Translation: The RNA(þ) is directly translated as a single endoplasmic reticulum (ER)-bound polyprotein. The 50 cap structure of the viral mRNA promotes assembly of eukaryotic initiation factors (eIFs) such as eIF4E and eIF4G and recruits ribosomes on the mRNA. In addition, in spite of the fact that the mRNA lacks a poly(A), the poly(A)-binding protein (PABP) interacts with the 30 UTR, suggesting that the viral genome is circularized though PABP–eIF4G interaction. The polyprotein is processed by viral and cellular proteases into three (then four) structural proteins and seven NS proteins (some of which are highlighted). (6) Replication: The NS proteins (highlighted) actively replicate the viral RNA(þ) in replication complexes (RCs) associated with cellular membranes, producing complementary RNA(), which in turn is used as a template to produce RNA(þ), that functions as genomic RNA. (7) Assembly: Following RNA replication and translation, virus assembly is achieved when one copy of RNA interacts with several copies of C protein forming NCs that are enveloped by the heterodimer PrM-E, to assemble into immature virus particles that bud into the lumen of the rough ER. (8, 9) Maturation: Virus particles transit through the Golgi (G) and the trans-Golgi network (TGN) where PrM is cleaved by cellular furin, resulting in the formation of particles containing the Pr, M, and E proteins. (10, 11) Exocytosis and release: The mature virus particle migrates to the cell membrane and is released from the cell as is also the Pr protein. N: nucleus.

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in the C-terminal region is believed to be the receptor recognition and binding domain and has an immunoglobulin-like fold (reviewed in Chin et al., 2007; Huerta et al., 2008). The interaction of the E protein with the receptor(s) leads to a series of events on the viral particle and in the cell membrane, and also in the cytoskeleton, allowing entry of the virus. Two types of cell receptors appear to be involved in facilitating entry of DENV into the human target cell, depending on the cell (reviewed in Huerta et al., 2008). The first type corresponds to receptors of low affinity and specificity, including aminoglycan-type adhesion molecules such as heparan sulfate that are expressed in many cell types (Germi et al., 2002; Lin et al., 2002). The second type corresponds to lectin-type receptors such as DC-SIGN (dendritic cell-specific intercellular adhesion molecule 3grabbing non-integrin) expressed in some antigen-presenting cells such as immature DCs (Lozach et al., 2005; Tassaneetrithep et al., 2003). These molecules presumably concentrate viral particles on the cell surface (Tassaneetrithep et al., 2003; Lozach et al., 2005; Reyes-del Valle et al., 2005; reviewed in Huerta et al., 2008) and facilitate recognition of the second type of receptors such as the heat- shock proteins (HSP) 70 and 90 (HSP70 and HSP90). This interaction induces the formation of endosomes; this phenomenon designated as ‘‘receptor-mediated endocytosis’’ (Fig. 2, step 2) allows virus entry into the cell (Reyes-del Valle et al., 2005). However, the implication of HSP70/90 as receptors is still a matter of debate and could depend on the cell types examined (Cabrera-Hernandez et al., 2007; Chavez-Salinas et al., 2008; Reyes-del Valle et al., 2005). Both these proteins are associated with microdomains, whose interaction is important for DENV entry. Interestingly, HSP70/90 has been postulated to be implicated in the entry of other viruses such as Hepatitis E virus (Zheng et al., 2010) and flaviviruses (Lin et al., 2007; Ren et al., 2007). Certain proteins known as adaptor proteins such as GRP78 are believed to also be part of the DENV entry complex, but their functions remain undefined (Cabrera-Hernandez et al., 2007). Associations between E and adaptor proteins are designated here as ‘‘fusion complexes.’’ The presence of lipid rafts or coated pits in virus fusion complexes indicates that vesicle formation depends on cholesterol (in human cells) or clathrin (in mosquito cells; Acosta et al., 2008; Lee et al., 2008a; van der Schaar et al., 2008). Together, these results show that regardless of the receptor used, at least two mechanisms of flavivirus entry into insect and mammalian cells exist: virus entry can occur by direct fusion of the virus to the cell membrane or by clathrin-dependent endocytosis (Chu and Ng, 2004; Chu et al., 2006; Hase et al., 1989a; Hase et al., 1989b; Krishnan et al., 2007; Lim and Ng, 1999; Se-Thoe et al., 2000; Suksanpaisan et al., 2009). Based on experiments in which it was shown that ablation of clathrinmediated endocytosis only reduces virus entry (Chu and Ng, 2004; Chu et al., 2006; Krishnan et al., 2007), another independent pathway of entry

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was proposed (Suksanpaisan et al., 2009). However, it should be stressed that different cell types were used in the various experiments referred to earlier. Another mode of entry for DENV is by ADE (see Section II.D; Kou et al., 2008). It has been demonstrated that infection of U937 monocytes is induced by ADE-mediated FcgRI, as is also infection of K562 leukaemic cells by FcgRII (Rodrigo et al., 2006). Although the role of FcgRIII in ADEDENV infection remains unclear, it was recently reported that the immune tyrosine activation motif of FcgRII is essential to mediate ADE (Moi et al., 2010). Interestingly, in mature DCs but not in immature DCs expressing high levels of DC-SIGN, ADE requires the Fcg receptor IIa (Boonnak et al., 2008). Finally, lipid rafts are necessary for AD-mediated infection of U937 cells by DENV (Puerta-Guardo et al., 2010).

B. Virus adsorption and fusion to the endosome membrane Evidence obtained to date suggests that the pathway of DENV entry after attachment to the host cell is primarily by clathrin-dependent endocytosis (Fig. 2, step 2). At pH 6.2–6.4, homodimer dissociation is facilitated and the resulting E monomers present a fusion loop that anchors to the endosome membrane, catalyzing the formation of E protein homotrimers (Heinz et al., 2004). This association induces a bend in the endosomal membrane promoting formation of a bridge (hemifusion stalk) between the virus and the endosomal membrane, leading to fusion. Thus, the nucleocapsid (NC) is released into the cytoplasm (Fig. 2, steps 3 and 4); how this occurs is largely unclear. After uncoating, the DENV genome is poised for translation by the host-cell machinery. Although the processes leading to fusion complex formation and to the E conformational changes involved in membrane fusion are established, no satisfactory agreement has been reached as to whether early and/or late endosomes are the site where viral fusion complexes form (Krishnan et al., 2007). Using HeLa cells, chemical inhibitors, and small interfering RNAs (siRNAs), it was demonstrated that Rab 5 (Rabaptin-5), a small GTPase protein that is a marker for early endosomes, showed that early endosome components are indispensable for virus amplification (Krishnan et al., 2007).

C. Intracellular transport of the viral genome Little is known about the mechanisms involved in the disassembly of the NCs, the transport of incoming genomes to the rough endoplamic reticulum (RER), or the recruitment of ribosomes by genome RNA. Intracellular transport of NCs/genomes is mediated by direct interaction with the cytoskeletal transport machinery (reviewed in Do¨hner and Sodeik, 2005;

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Kanlaya et al., 2009). The cytoskeleton is composed of actin and vimentin filaments, microtubules (MTs), and intermediate filaments (IFs) that are constantly modified during the cell cycle. These modifications include morphological changes and activation, that is, maturation and rearrangements that can be enhanced by the viral pathogen (Chen et al., 2008). DENV can alter the components of the cytoskeleton, but how this is achieved is poorly understood; it requires actin-rich regions endowed with a dynamic reorganization mechanism (Chen et al., 2008; Talavera et al., 2004) and also affects vimentin filaments, MTs, and IFs. In the presence of DENV, the arrangement is modified and the vimentin filaments retract from the cell periphery and from around the nucleus (Chen et al., 2008). Other studies showed that infection of human microvascular endothelial cells-1 (HMEC-1) with DENV-2 promotes actin reorganization, regulated by the GTPase proteins, ras-related c3 botulinum toxin substrate 1 (Rac1), and cell division cycle 42 (Cdc42); this is critical for the formation and function of filopodia, necessary for viral entry (ZamudioMeza et al., 2009). Thus, the cytoskeletal transport machinery may facilitate the transfer of the viral genome to the ribosomes. Yet, the mechanism involved and the time necessary for capsid disassembly and RNA release are unknown.

D. Genome expression Liberated from its structural proteins, the DENV genome (Fig. 1) becomes accessible to perform its three major functions: (1) as mRNA for the synthesis of the viral polyprotein (translation), (2) as template for the synthesis of further RNA strands (replication), and (3) as genome incorporated into new viral particles (encapsidation). It is important to bear in mind that DENV genome translation is coupled to replication, since the viral genome must be translated to synthesize the NS proteins required for RNA replication (reviewed in Harris et al., 2006). Translation of the DENV RNA(þ) produces a single endoplasmic reticulum (ER)-bound polyprotein (Fig. 2, step 5). Synthesis of the viral polyprotein is initiated at the 50 proximal AUG codon in the RNA. The cap structure of the viral mRNA facilitates assembly of eukaryotic initiation factors (eIFs) such as eIF4E and eIF4G and recruits ribosomes on the mRNA. Interestingly, the poly(A)-binding protein (PABP) can bind to the 30 UTR (Polacek et al., 2009b) even though the genome lacks a poly (A) tail, suggesting that circularization of the genome via interaction with the PABP-eIF4G complex could be required for efficient translation (Fig. 2, step 5). Interestingly, initiation of DENV mRNA translation can also occur efficiently when cap-dependent translation is inhibited, such as when the cap-binding protein eIF4E is limiting (Edgil et al., 2006). Experiments

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performed in the presence of inhibitors of cellular-capped mRNA translation, showed that DENV RNA or a reporter mRNA flanked by the DENV 50 UTR and 30 UTR was efficiently translated. Translation by a cap-independent initiation mechanism may facilitate amplification of viruses that multiply in divergent hosts (e.g., mosquitoes and humans) and may also help the virus accommodate to various intracellular environments. The highly conserved 3’SL structure present in the 3’UTR (Fig. 1B) facilitates mRNA binding to polysomes and is a major player in promoting efficient DENV-2 mRNA translation (Holden and Harris, 2004); during the first round of translation, it stimulates translation in the absence of viral proteins. Natural variants of DENV-2 that differ in their ability to multiply in primary human cells appear to vary in their ability to synthesize viral proteins in vivo and in vitro (Edgil et al., 2003). These differences are linked to mutations in the 3’UTR and suggest that other regions than the coding sequence are important for efficient translation and could be activated by modifying either RNA–RNA or RNA–protein interactions. Certain cell proteins bind to the 3’UTR. Indeed, in addition to certain translation initiation factors important for cap-dependent DENV RNA translation, the 3’SL binds several cell proteins such as the eukaryotic elongation factor (eEF) 1A enhancing viral RNA transcription (Davis et al., 2007). Other reports have shown that the La autoantigen, the polypyrimidinetract-binding protein, and the Y-box-binding protein-1 bound to the 3’UTR (Davis et al., 2007; De Nova-Ocampo et al., 2002; Garcia-Montalvo et al., 2004; Paranjape and Harris, 2007), but the functional roles of these interactions in replication and/or translation have not been clearly established. Some results have proposed that translation and subsequent replication occur within intracellular membranes structures, whose origins are not well defined (reviewed in Clyde et al., 2006; Miller and Krijnse-Locker, 2008).

E. Genome replication The association of the machinery for DENV negative-sense RNA [RNA ()] synthesis with host intracellular membranes is poorly understood, but the membranes probably play a structural and organizational role in forming the replication complex (RC). However, it is known that following translation, viral NS proteins together with host factors form the RC responsible for the synthesis of RNA() complementary to the RNA(þ), resulting in an intermediate double-stranded (ds) RNA. The complete DENV RNA replication cycle includes the synthesis of RNA() and of new RNA(þ) that functions as mRNA (Fig. 2, step 6), and as genomic RNA to form new virus progeny. RNA synthesis is regulated by

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sequences located in the 50 and 30 UTRs: the 50 –30 CS and 50 –30 UAR sequences (Fig. 1A and B) mediate Mg2þ-dependent and cell proteindependent circularization, resulting in long-range RNA–RNA interactions important for DENV RNA replication (Alvarez et al., 2005a,b; Khromykh et al., 2001; Lo et al., 2003). Circularization is further facilitated by the presence of a 6-nt-long 50 DAR sequence (Friebe and Harris, 2010). Although the specific role of NS3 in replication remains unclear, its helicase domain could be involved in unwinding RNA secondary structures present in the 30 UTR and assisting the first steps of replication. NS3 is a multifunctional protein with RNA-stimulated nucleoside triphosphatase (NTPase), ATPase/helicase, and RTPase (Sampath et al., 2006; Wang, et al., 2009a; Yon et al., 2005) activities that are essential for viral RNA replication and capping. SLA at the 50 end of the 50 UTR is the promoter for the RdRp activity of NS5 and RNA synthesis (Filomatori et al., 2006). In transfected cells, NS5 interacts with the 50 SLA, but surprisingly not with the 30 UTR (Filomatori et al., 2006). Various structures in the 50 SLA (UAR, cHP, and CS) are involved in circularization of the DENV genome and in promotion of RNA synthesis (Friebe and Harris, 2010; Lodeiro et al., 2009). Despite sequence and structural differences between the 50 SLAs of DENV-1 and DENV-2, the RdRp of DENV-2 efficiently uses the 50 SLA of DENV-1 for RNA replication (Filomatori et al., 2006; Hahn et al., 1987; Yu et al., 2008b). Thus, the requirement for the homologous 30 SL for viral replication is related to specificities of other viral NS proteins in addition to NS5 which not only functions as the RdRp, but also acts as the MTase implicated in capping of the viral RNA. Alternatively, the specificity of NS5 could be altered by its interaction with other NS proteins or with cell proteins. A single mutation within the 50 UAR can decrease RNA synthesis up to 3000 fold during DENV genome replication (Alvarez et al., 2008). In addition, genomes with disruption of the stem of SLB by mutations that still maintain complementarity with the 30 UAR, can be translated and replicated. Therefore, the 50 UAR sequence acts as a cis-element regulating DENV RNA replication. In the DENV genome, interaction between the 30 and 50 CSs also facilitates hybridization of the 50 –30 UARs that form a 15-ntlong base-paired structure interrupted by a C bulge and a G–G mismatch (Fig. 1B; Alvarez et al., 2008). Although hybridization between the 50 and 30 UARs requires interaction between the 50 and 30 CSs, interaction between the 50 and 30 CSs does not depend on interaction between the 50 and 30 UARs (Polacek et al., 2009a). Studies performed in vitro using viral subgenomic (sg) RNA templates containing the 50 and 30 CSs showed that interaction between the two terminal regions is necessary for the synthesis of RNA() from RNA(þ) (Ackermann and Padmanabhan, 2001; Nomaguchi et al., 2004; You et al., 2001). Recently, the presence of a small sgRNA derived from the 30 UTR by cell nuclease digestion and

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essential for virus-induced pathogenicity, was identified in DENVinfected cell cultures and in animal tissues (Liu et al., 2010; Pijlman et al., 2008). Both the sgRNA and genomic RNA accumulated together during viral RNA replication. It will be interesting to define the function of this noncoding sgRNA in the virus life cycle. The 30 UTR() could act as a promoter element for the synthesis of new RNA(þ) as does the 30 UTR(þ) for the synthesis of RNA(). In addition, the 30 UTR() of DENV interacts with La, a nuclear protein that participates in cell mRNA transcription initiation, forming a ribonucleoprotein complex in association with other cell proteins such as calreticulin and the protein disulfide isomerase (Yocupicio-Monroy et al., 2003). Circularization of the viral RNA favors coupling of translation and replication. This occurs in the cytoplasm on induced cell membranous structures probably derived from the ER, and in vacuoles induced by the virus and known as viral RCs. NS2A, in addition to playing a direct role in viral RNA replication, could also be the viral protein that targets the viral RCs to membrane organelles (Mackenzie et al., 1998). NS4B which presents at its C-terminus a highly hydrophobic fragment (designated 2K) that functions as a signal sequence, serves for the translocation of NS4B into the ER lumen, where it is a component of RCs (Miller et al., 2006). In the membrane-bound viral RCs, NS4B is associated with NS3 and dsRNA, a marker for replicating viral RNA. During this process, proteolytic removal of a 2 kDa peptide from NS4A is important for the induction of membrane alterations that may harbor the viral RCs (Miller et al., 2007). Dissociation of NS3 from ssRNA by NS4B in vitro enhances the NS3 helicase activity, suggesting that NS4B can modulate DENV replication (Umareddy et al., 2006). Viral genome replication takes place in the RCs (reviewed in Bartenschlager and Miller 2008), although some replication activity has been detected in the nucleus (Uchil et al., 2006). The question, however, remains how the RCs are formed. A role of autophagy in virus replication has recently become a new emerging field of investigation. Autophagy develops in an area of the cytoplasm where a membrane preexists and expands the membrane to form doublemembrane vesicles; it is part of a lysosomal degradation pathway that is important for cell remodeling and development, and that is also involved in various disease processes. DENV appears to also induce autophagy as a means of enhancing replication in the cell (Colombo, 2005; Lee et al., 2008b), and viral RNA replication has been associated with double-membrane structures (Uchil and Satchidanandam, 2003; reviewed in Miller and Krijnse-Locker, 2008) that are hallmarks of autophagosomes (Dunn, 1990). Other studies showed that the induction of autophagy by DENV-2 and DENV-3 resulted in an increase in the titers of extracellular and intracellular virus (Khakpoor et al., 2009). Moreover, DENV-2 triggers autophagy by increasing the expression of LC3-II (light chain 3 form II of the

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microtubule-associated protein 1) and green fluorescent protein (GFP)-LC3 dot formation. In addition, LC3-II is associated with autophagosome membranes and colocalizes with viral dsRNA, NS1, and the ribosomal protein L28 (Kuma et al., 2004; Panyasrivanit et al., 2009b). Inhibition of fusion of autophagosomes and amphisomes with lysosomes decreases DENV-3 production, suggesting a role for autophagosomes in the DENV life cycle (Panyasrivanit et al., 2009a). Based on results showing that DENV components colocalize with markers of autophagic and endosomal vesicles, it was proposed that amphisomes constitute a site of DENV genome replication and translation. The activation of the cell autophagy machinery to promote DENV RNA replication depends on the ATG5 protein, an initiator of autophagosome formation during autophagy progression (Kuma et al., 2004; Lee et al., 2008b). Inhibition of autophagy by ATG5 knockout or treatment of cells with lysosomes that inhibit fusion of autophagosomes and amphisomes, is associated with a decrease in DENV replication and reduced progeny virus particle formation (Khakpoor et al., 2009; Limjindaporn et al., 2009). Interestingly, there appears to be a link between DENV entry and replication/ translation in terms of a continued association of the virus life cycle with membranes of an endosomal–autophagosomal lineage (Panyasrivanit et al., 2009a). Thus, DENV replication/translation is coordinated by cis-acting genomic sequences, viral proteins, and cell factors. Future studies should clarify how autophagic membranes and DENV coordinate the autophagic processes.

F. Maturation and release of DENV from the host cell After polyprotein synthesis and genome replication have occurred, the next step leading to viral particle formation is encapsidation. Although the first indications of interaction between the viral RNA and C were demonstrated over a decade ago (Khromykh et al., 1999), the precise mechanisms involved in RNA-C complex (or NC) formation are unclear. Recently, Samsa et al. (2009) reported that mature C protein is associated with lipid droplets (LDs), defined as ER-derived organelles located in the cytoplasm and that contain various proteins surrounded by a core of neutral lipids and a monolayer of phospholipids; this association is triggered by hydrophobic residues present in the center of C (Ma et al., 2004). Infection of cells by DENV stimulates the formation of LDs, and decreasing the number of LDs reduces virus replication. Strong evidence suggests that viral RNA-C protein assembly occurs in membrane structures derived from the ER (Fig. 2, step 7), forming NCs. In addition to its RNA-binding motifs, the N-terminal region (aa 21–100) of the C protein contains four hydrophobic a helices essential for association with the ER (Ma et al., 2004). Thus, the partially assembled virus NCs bud from the lumen of the ER (Fig. 2, step 7) and become enveloped by a lipid

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membrane that contains, in addition to NCs, the two structural proteins, prM and E, in the form of prM-E heterodimers (Mukhopadhyay et al., 2005, Wang et al., 2009b) that are incorporated into immature virions by interaction with the NC; this step constitutes the assembly of immature viral particle. Yet, NCs are not prerequisites for the formation and secretion of viral particles, since sub-viral particles composed only of prM and E are also secreted from DENV-infected cells (Mukhopadhyay et al., 2005). The E protein undergoes reversible pH-dependent conformational changes during egress of virions through the secretory pathway. In the immature state, the prM-E heterodimers are grouped as trimers, whereas in the mature particles, there are E homodimers, and E forms homotrimers when the virus fuses with the host cell endosomal membrane (Bressanelli et al., 2004). During this transport process, the virus particles pass through the TGN where the prM protein is cleaved by the protease furin, but the pr segment continues to be associated with the virion until the virion is released from the cell. The retention of the pr segment prevents premature binding of the E protein to the exosomal membrane during transit of the virus in the acidic environment of the TGN. These mature viral particles are released from the TGN into the cytoplasm and are transported to the outer cell membrane by exocytosis (Fig. 2, step 10; Yu et al., 2008a, 2009; reviewed in Perera and Kuhn, 2008). Finally, the cleaved pr and virions are released into the extracellular medium upon particle secretion (Fig. 2, step 11; Yu et al., 2009). Virus maturation in the lumen of the TGN produces mature virions composed of one copy of RNA, 90 homodimers of E, and 180 copies of prM. The transition from immature (spiky) to mature (smooth) particles due to the cleavage of prM occurs during transit of the particles through the Golgi into the TGN (Fig. 2, steps 8 and 9; Wang et al., 2009b). Cleavage of prM to M by furin is essential for DENV infectivity (Zybert et al., 2008). Indeed, treatment of immature viruses with exogenous furin restores viral infectivity. Analyses performed with other flaviviruses suggest that prM cleavage is required for full maturity and infectivity of the virus particles (Wengler and Wengler 1989). However, cleavage of DENV prM is inefficient, as various cell types infected with DENV-2 release large amounts of virions with unprocessed prM (Elshuber et al., 2003). In addition to preventing fusion of E to the exosomal membrane (Yu et al., 2008a), pr retention could also (1) constitute a mechanism favoring flavivirus trafficking and stability in the cell secretory pathway, (2) be required for interaction with the vacuolar-ATPases (V-ATPases) whose silencing leads to reduction in DENV replication, and (3) establish a suitable pH environment for efficient virus secretion (Duan et al., 2008). As mentioned earlier, acidification of intracellular organelles such as components of the secretory pathway is crucial for infectivity (Yu et al., 2008a, 2009). Previous reports have shown that V-ATPases are

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multisubunit enzymes that acidify various organelles, including lysosomes and components of the secretory pathway, facilitating protein processing and acid-dependent protein degradation during DENV infection (Duan et al., 2008). Residues 76–80 of prM participate in the interaction with V-ATPases, and the V-ATPase-virus interaction is critical for efficient virus entry and egress. Thus, the viral particles assemble and bud from the lumen of the ER, and virus maturation occurs in the TGN; finally, the mature virus particles are released by exocytosis. In addition, it was recently demonstrated that the human immunoglobulin heavy chain-binding proteins, calnexin and calreticulin, all ER-resident chaperones, interact with the E protein (Limjindaporn et al., 2009). Silencing of the three corresponding genes using siRNAs affected the production of infectious DENV, suggesting that these chaperones participate in the folding and assembly of the viral proteins. The E protein of DENV is glycosylated in the residues 67 and 153/154. Loss of either of the two E protein glycans enhanced infectivity of variants for mosquito cells and reduced virus release in mammalian cells. (Hanna et al., 2005; Lee et al., 2010).

V. SIRNAS AND NEW STRATEGIES TO CONTROL DENV REPLICATION An understanding of the cellular components involved in DENV infection is imperative, as it should allow better comprehension of dengue pathogenesis and lead to the elaboration of gene-targeted inhibitor strategies. However, only a few reports of inhibitory factors of DENV infection have appeared. Lately, siRNAs have been used in vivo as a gene-silencing approach to decrease DENV replication. dsRNA is cleaved by Dicer, an RNAse-III-type enzyme, into 21–25-nt-long siRNAs. siRNAs have been used as a knockdown mechanism to study cellular and viral genes involved in virus replication and constitute a new pathway for clinical treatment of various infectious diseases. Several reports have demonstrated the efficiency of the siRNA strategy to block DENV production (Ang et al., 2010; Mukherjee and Hanley, 2010; Subramanya et al., 2010). The nucleotide sequence of the 30 UTR which is common to the genome of all four DENV serotypes, was used to design siRNAs, and these siRNAs silenced DENV RNA replication. Adeno-associated virus encoding siRNAs targeted to the 30 UTR of the DENV RNA also reduced DENV infection in Vero cells and in human DCs in a dose-dependent manner (Zhang et al., 2004). Moreover, infection of human dermal microvascular endothelial line-1 cells with DENV-2 induces high expression of b3 integrin, and preincubation of the virus with soluble ab integrin or silencing of the b3 integrin gene using siRNAs

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drastically reduces virus entry (Zhang et al., 2007). In Huh7 cells, the use of siRNAs targeting genes implicated in clathrin-mediated endocytosis inhibited DENV entry into target cells (Ang et al., 2010). In HeLa cells, silencing of the gene encoding the V-ATPase, a proton pump that is key to establishing the low pH of endosomal compartments resulted in decreased DENV replication (Krishnan, et al., 2007), demonstrating the requirement of V-ATPase in endosomal DENV entry. Moreover, knockdown of the Rab 5 GTPase gene by siRNAs revealed that DENV infection requires the expression of Rab 5, a key regulator of transport to early endosomes. Cholesterol is an important element in DENV infection; its depletion with methyl-bcyclodextrin or its chelation with filipin III, a member of the polyene family of antibiotics, diminishes entry of DENV or Japanese encephalitis virus (Lee et al., 2008a). Furthermore, selective silencing of enzymes involved in cholesterol biosynthesis using siRNAs inhibits DENV replication in A549 cells (Rothwell et al., 2009). Both studies suggest that cholesterol modulation affects DENV replication, although the mechanism(s) whereby this occurs is as yet unknown. DENV replication occurs in association with the ER, where the viral particles assemble; in a search for the role of host ER chaperones in these processes, it was demonstrated that siRNAs that separately silence three ER-resident chaperones, the immunoglobulin heavy chain-binding protein known as BiP, calnexin, or calreticulin, significantly decreased the yield of infectious virus production (Limjindaporn et al., 2009). Moreover, using image-based immunofluorescence assays, inhibitors of the c-Src protein kinase were shown to be potent inhibitors of DENV replication (Chu and Yang, 2007), preventing virus assembly within the RC. The data suggest that siRNAs could be used to hinder DENV infection and could serve as a therapeutic strategy. Interestingly, infection by DENV-2 of cultured A. aegypti cells or DENV-2 administered orally to adult mosquitoes, led to the production of siRNAs specific for the virus, whereas impairing the siRNA machinery increased virus production in this vector (Sa´nchez-Vargas et al., 2009). Infection of Drosophila melanogaster S2 cells with any of the four serotypes of DENV induced siRNA production (Mukherjee and Hanley, 2010). Knockdown of one gene implicated in the RNA interference (RNAi) machinery led to enhancement of viral replication. Consequently, infection of insect cells activates an antiviral response mediated by siRNAs.

VI. CONCLUSIONS In recent years, new approaches such as transcriptomic/proteomic and microarray techniques to study the integral aspects of virus life cycles have led to more profound knowledge concerning the relationship between virus and host. It was known that formation of infectious

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DENV requires the concerted action of viral structural proteins. More recently, the importance for viral genome expression of viral NS proteins and cell proteins, and of regions of the viral genome that lie within the ORF are being recognized, as is also the coupling of viral RNA replication and virus assembly. These approaches have also demonstrated that NS proteins participate in the localization of active RCs and mediate formation of specialized assembly sites to regulate the release of RNA from the RCs and RNA packaging in the budding virus. Although replication occurs in the cytoplasm, NS5 has also been immunolocalized in the nucleus where its specific role in the virus life cycle appears to be to antagonize the antiviral response. However, the functions of other DENV proteins that traffic between the nucleus and cytoplasm are unclear. Likewise, the function of the C protein in the nucleus is unknown. These observations nevertheless illustrate the essential roles of the viral proteins and their use as potential targets for genetic therapeutic strategies against DENV infection. The identification of host cell factors important for DENV infection remains an essential but largely unexplored area. Although the functions of the NS proteins in the production of infectious progeny are beginning to emerge, further studies should aim at elucidating their specific mechanisms of action so as to exploit this knowledge in the development of highly efficient antiviral drugs for dengue treatment.

ACKNOWLEDGMENTS Thanks are due to Diana Giraldo and Juan Guillermo Betancur for their help with the figures. We are very grateful to Margo Brinton for her constructive and valuable comments regarding the manuscript. We regret that we have not been able to refer to all the published works that have recently promoted the expansion for this area of research. This study was supported by the Universidad de Antioquia, Colciencias, the Institut Jacques Monod, and by grant 111540820517 from Colciencias. CP was supported by a Fellowship from the FriederischMiescher Institute/Novartis and by grant 111534319145 from Colciencias; ST is grateful to Colciencias for a Ph. D. fellowship.

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CHAPTER

2 Evolution of Human Papillomavirus Carcinogenicity Koenraad Van Doorslaer* and Robert D. Burk*,†,‡,§

Contents

Abstract

I. Papillomavirus Classification II. Clinical Implications of Human Papillomavirus Infections III. HPV and Cervical Carcinogenesis IV. HPV Life Cycle V. HPV Viral Carcinogenicity is Correlated with the Early Viral Region VI. The E6 and E7 Viral Oncogenes VII. Evolutionary History of the E6 and E7 Oncoproteins VIII. Toward a Biochemical Understanding of Alphapapillomavirus Oncogenicity IX. Conclusions and Perspectives Acknowledgments References

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Members of the Alphapapillomavirus genus are the causative agent for virtually all cases of cervical cancer. However, strains (commonly referred to as types) within this genus span the entire range of pathogenicity from highly carcinogenic (e.g., HPV16, odds ratio ¼ 281.9, responsible for 50% of all cervical cancers), moderately

* Department of Microbiology and Immunology, Albert Einstein Cancer Center, Albert Einstein College of {

{

}

Medicine, New York, USA Department of Pediatrics, Division of Genetics, Albert Einstein Cancer Center, Albert Einstein College of Medicine, New York, USA Department of Epidemiology and Population Health, Albert Einstein Cancer Center, Albert Einstein College of Medicine, New York, USA Department Obstetrics, Gynecology and Woman’s Health, Albert Einstein Cancer Center, Albert Einstein College of Medicine, New York, USA

Advances in Virus Research, Volume 77 ISSN 0065-3527, DOI: 10.1016/S0065-3527(10)77002-0

#

2010 Elsevier Inc. All rights reserved.

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Koenraad Van Doorslaer and Robert D. Burk

carcinogenic (e.g., HPV31) to not carcinogenic (e.g., HPV71). The persistent expression of the viral oncoproteins (E6 and E7) from HPV16 has been shown to be necessary and sufficient to transform primary human keratinocytes in vitro. A plethora of functions have been described for both oncoproteins, and through functional comparisons between HPV16 and HPV6, a subset of these functions have been suggested to be oncogenic. However, extrapolating functional differences from these comparisons is unlikely to tease apart the fine details. In this review, we argue that a thorough understanding of the molecular mechanisms differentiating oncogenic from nononcogenic types should be obtained by performing functional assays in an evolutionary and epidemiological framework. We continue by interpreting some recent results using this paradigm and end by suggesting directions for future inquiries.

I. PAPILLOMAVIRUS CLASSIFICATION In 2004, de Villiers and colleagues proposed quantitative thresholds for classification of PVs based on sequence identity, topology of phylogenetic trees, and biological characteristics (de Villiers et al., 2004). Papillomaviruses sharing more than 60% nucleotide sequence identity across the major structural gene, L1, were classified within the same genus. These genera were named according to the Greek alphabet (de Villiers et al., 2004). This was the first viral classification system based on sequence similarity recognized by the ICTV (Fauquet et al., 2005). Within a specific genus, distinct PV strains, commonly referred to as ‘‘types’’, were described as a virus for which the entire genome was cloned, and the L1 ORF shared less than 90% sequence similarity with any other previously defined type. Additional research since 2004 has resulted in the identification of PVs from a large number of different host species ranging from turtles, birds, and mammals. A recent publication updating papillomavirus nomenclature assigned the 189 known PVs to 29 genera (Bernard et al., 2010). Figure 1 shows the phylogenetic relationships between these types based on L1 the ORF.

II. CLINICAL IMPLICATIONS OF HUMAN PAPILLOMAVIRUS INFECTIONS The phylogenetic tree shown in Fig. 1, clusters most HPVs into three main clades (a, b, and g), roughly corresponding to preferred tissue tropism. In the remainder of this review, we will focus on the a-PV genus, but for completeness, we provide a quick overview of the other two main HPV genera.

Evolution of Human Papillomavirus Carcinogenicity

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Alpha a12 a11

a13 a1

a2 a7

a9

Omega Dyodelta

a5 a8

Lambda

a6 a10

Kappa

l4 l2 l3

l1

Sigma

a4

Mu

Nu

k2 k1

Iota

m1

a3

m2

d4

a14

d5

Dyotheta

Delta

d2

Psi Dyozeta Theta Eta Dyoepsilon

d1 d3

b5 b4 b3

c2 c1

b6

Rho Phi Omikron

Xi

p2 n2 p1

n1

b2 g7

b1

g10

g4 g2 g3

Upsilon

g6

Pi

g8

g9

Dyoeta

Chi

g5

g1

Beta

Epsilon Zeta Dyoiota

Tau

Gamma

FIGURE 1 Bayesian phylogenetic tree based on the L1 nucleotide sequences of 189 papillomaviruses. Adapted from Bernard and colleagues (Bernard et al., 2010).

The types within the b-PV genus present as cutaneous infections that are mostly latent in the general population (de Villiers et al., 2004); however, in a subset of immunocompromised patients, these viral types can cause epidermodysplasia verruciformis (EV) (Dell’Oste et al., 2009). These patients often develop cutaneous squamous-cell carcinoma. In 90% of the cases, these lesions can be shown to be positive for HPV-5 or -8 (Orth, 1986). However, at present, there is insufficient evidence to consider HPV-5 or -8 as independent human carcinogens (Bouvard et al., 2010). The members of the g-PV genus present as mainly commensal infections of the skin (de Villiers et al., 2004). Interestingly, whereas most b- and gPVs have been isolated from the skin, recent evidence also suggests high prevalence of these viral types in the oral cavity (Burk, manuscript in preparation). The a-PV genus contains viruses infecting cutaneous and mucosal keratinocytes. Typical cutaneous lesions present as verruca plantaris, verruca plana, or ‘‘butchers warts’’ depending on the infecting HPV type. Infections with HPV6 and 11 are mainly associated with condylomata accuminata; however, they are also the main culprits in the etiology of laryngeal papillomas and recurrent respiratory papillomatosis.

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Within the a-PV genus, most research has focused on cervical carcinogenesis (see Section III); however, infection with HPV16 is responsible for the preponderance of HPV-induced tumors at sites other than the cervix. HPV16 has been found to be responsible for the majority of vaginal, vulvar, penile, and anal cancers (Munoz et al., 2006). In addition, increasing evidence is linking a-HPV infection to the development of head and neck squamous-cell carcinoma (McKaig et al., 1998), and conjunctival cancers (Nakamura et al., 1997).

III. HPV AND CERVICAL CARCINOGENESIS Yearly, the worldwide incidence of cervical cancer has been about 510,000 new cases, with nearly 300,000 patients dying (WHO: http://www.who. int/vaccine_research/diseases/hpv/en/). Most cases of cervix cancer (80%) occur in developing countries, particularly Asia. Increasing epidemiological evidence has identified persistent infection with oncogenic human a-papillomaviruses as the causative agents of cervical carcinoma. However, of the over 120 identified HPVs, only a limited subset of viruses (about 20 so-called oncogenic types (OTs)) have carcinogenic potential (Bouvard et al., 2010), with HPV types 16, 18, 31, 33, 45, 52, and 58 showing the highest association with cervical malignancies (Smith et al., 2007). Infections are mainly transmitted through direct skin or mucosal contact, with a substantial probability of transmission with every sexual contact (Burchell et al., 2006). There appears to be a step-wise progression from infection through progressively severe intraepithelial neoplasia (from CIN stage 1 to CIN3) to invasive cancer. However, at each stage there is the possibility for viral clearance and/or regression of the lesion (Schiffman and Kjaer, 2003; Fig. 2). So, even though approximately 45% of women will acquire an infection within the first 36 months of sexual activity (Ho et al., 1998), most infections will be cleared within 2 years. This is consistent with Normal Cervix

Infection Clearance

Infected cervix

Persistence

Invasion CIN3

Cancer

Regression

Mild cytological abnormalities

FIGURE 2 Model of HPV infection of the cervix natural history and progression to cancer. Most infections are quickly cleared through the innate immune system. Even following establishment, the host immune response can still clear the viral infection. The persistently infected epithelium (1–10 years) will support clonal progression, leading to CIN3. In a final, not well-understood step, the lesion progresses into invasive carcinoma.

Evolution of Human Papillomavirus Carcinogenicity

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the data that  90% of new infections with carcinogenic HPV types will not be of clinical significance (Rodriguez et al., 2010). Nevertheless, even though persistent OT HPV infection and malignant progression are highly associated (Ho et al., 1995; Nobbenhuis et al., 1999; Schlecht et al., 2001), it is the potential to progress to malignant lesions that separates the OT viruses from equally well-persisting NOT viruses (Burk et al., 2009; Schiffman et al., 2005). This observation suggests a functional difference between carcinogenesis independent of immune evasion and persistence.

IV. HPV LIFE CYCLE HPV is a circular double-stranded DNA virus with a genome of about 7900 bp. The genome codes for eight proteins on the same sense strand and carries one upstream regulatory region (URR), also known as the noncoding region (Fig. 3). The genes are named based on their expression during epithelial differentiation (early (E) or late (L)). The viral life cycle is tightly linked to the differentiation state of the host cell it infects (Fig. 4). Following establishment of viral infection in the basal layer of the epithelium, the viral genome is maintained at about 100 copies per cell (Watts et al., 1983). URR

E6

E7

7,908/1

L2 5931

HPV16

1977

E1

3954

L1

E4 E5

E2

FIGURE 3 The structure and organization of the HPV genome. The reference HPV16 genome is shown. The PV genome consists of a circular double-stranded DNA genome. The genome is divided into three regions. We indicated the upstream regulatory region (URR, gray box). The viral genome carries all ORFs on a single sense strand, with the E4 ORF overlapping the E2 ORF. The early genes (E, solid arrows) and late (L, dashed arrows) structural genes are indicated.

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Koenraad Van Doorslaer and Robert D. Burk

Virus assembly/ virus release

L1

L2

E4

Virus release

Granular

Genome amplification

Viral DNA

Suprabasal

E1, E2, E4, E5

E6, E7

Epidermis Genome maintenance/ cell proliferation

Genome maintenance

Basal Dermis Mucosal

FIGURE 4 Epithelial infection by papillomaviruses. The key events that occur following infection are tightly linked to the differentiation state of the infected cell. The arrows next to the figure indicate the temporal pattern of the indicated proteins’ expression, while the color represents the amounts expressed. On the far right, the effects of timed expression are summarized. Cells with red nuclei are progressing through the cell cycle. Since cells above the basal layer are terminally differentiated, this implies that the viral oncogenes, E6 and E7, are subverting cell cycle control mechanisms. Epithelial differentiation activates the differentiation-dependent promoters, resulting in increased expression of the viral replication proteins (E1, E2, E4, and E5) and genome replication (green cells with red nuclei). While continuing on their differentiating path, cells containing amplified viral DNA (yellow) begin expressing the structural L1 and L2 genes. The green/yellow cells contain infectious particles, which are shed at the epithelial surface. It appears that increased expression of the E4 protein near the epithelial surface is involved in viral shedding. Figure adapted from Doorbar (2006).

The expression of the E1 and E2 proteins is thought to be essential for viral episomal maintenance. The E2 protein binds to a palindromic motif [AACCg(N4)cGGTT] in the URR (Blakaj et al., 2009), which allows for the recruitment of the E1 helicase (consensus motif AACNAT), which in turn recruits host replication factors to the viral origin of replication (Yang et al., 1991). The requirement of host factors (e.g., DNA polymerase a/primase (Bonne-Andrea et al., 1995; Conger et al., 1999; Masterson et al., 1998) and the single-stranded-DNA-binding protein RPA (Han et al., 1999)) for viral DNA replication implies that the terminally differentiated host cells need to reenter the cell cycle. The expression of two viral proteins, E6 and E7, has been shown to play a key role in this process (see Section VI). In order to achieve a productive infection, the viral genomic copy number needs to be amplified prior to being packaged into viral capsids.

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The event triggering productive infection is not well understood, but is related to the differentiation state of the host cell. It has been shown that a viral differentiation-dependent promoter element is key to the increased expression of the E1, E2, E4, and E5 proteins (Bodily and Meyers, 2005; Spink and Laimins, 2005). Following the beginning of genomic amplification, L1 and L2 expression is upregulated both at the RNA and protein levels (Schwartz, 2000). It is interesting that the L1 and L2 levels appear to also be controlled through the use of suboptimal codons (Zhou et al., 1999). Although the expression of L1 is sufficient to achieve viral packaging, the presence of a L2 protein substantially increases the efficiency (Zhao et al., 1998). In addition, it has been suggested that the E2 protein, through its specific binding to the viral genome, also plays a role in packaging (Zhao et al., 2000). Finally, expression of the E4 protein disrupts the integrity of the cornified cell layer assisting in efficient escape from the epithelial cell surface (Doorbar et al., 1991).

V. HPV VIRAL CARCINOGENICITY IS CORRELATED WITH THE EARLY VIRAL REGION Taxonomic classification of PVs places most HPVs in three main genera, roughly corresponding to body sites of infection. Essentially, all known HPVs infecting the cervicovaginal epithelium are contained within the a-PV genus (Fig. 1; Bernard et al., 2010; de Villiers et al., 2004). Consistent with this observation, the a-PV genus contains all the viruses for which there is sufficient epidemiological evidence to be considered oncogenic (Bouvard et al., 2010; Schiffman et al., 2005). The a-PV genus consists of three main clades, the high-risk (HR), which contains all the OTs, and two low-risk (LR1 and LR2) clades (Schiffman et al., 2005). However, previous work identified significant phylogenetic incongruence at the putative high-risk (i.e., cancer-associated, HR) node (Fig. 5; Narechania et al., 2005). A common oncogenic ancestor shared homology across the early gene ORFs, but not the structural (late) genes, that is, in the late gene trees, the HR- and LR-clades are mixed, whereas these clades are monophyletic and separate when considering the early genes. In addition, Fig. 5 shows two phylogenetic trees based on the E6 and E7 oncogenes; when comparing these trees, it can be noticed that both LR-clades form a monophyletic group, independent from the HR-clade. These data support the notion that HPV species groups followed distinct evolutionary paths, with differences noted for the early and late genes (Narechania et al., 2005). This might help explain how specific HPVs evolved to infect different cell types while preserving oncogenic capabilities (e.g., HPV16 and -18 are disproportionately associated with squamous and adenosquamous carcinoma, respectively; Smith et al., 2007).

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E1

E2 6 5

9 11

7

7

9 11 10 8 1

5 6 1 10 8 HPV54

HPV54

E6

3

3

15 4

4 15

2

2

1 10 8

E7

HPV54

L2

1 10 8

3

3

4 15

4 15 HPV54

2

2

6 5 7

9 11 7

9 11

6 5

15 3 4 10

L1

1 10 8 11 9

HPV54

HPV54

11 9

3

8

15 4 6

7 6 5 2

7 5 2

FIGURE 5 Phylogenetic trees based on the six main viral ORFs show the incongruence between each viral region. Maximum Likelihood phylogenentic trees were built on the nucleotide sequences of all-known HPVs within the a-PV genus. The optimal tree was selected from 50 independent ML searches under the GTRGAMMA model, as implemented in RAxML (Stamatakis et al., 2005). For improved readability, the branches were collapsed according to the accepted HPV classification (Bernard et al., 2010; de Villiers et al., 2004). Numbers next to the collapsed clade correspond to the species within the a-PV genus. HPV54 is the sole representative of the alpha-13 species. Branches are colored to illustrate viral risk classification; HR (red), LR1 (blue), and LR2 (green) (Burk et al., 2009). To test for incongruency between the different trees, the Shimodaira– Hasegawa test for topology (Shimodaira and Hasegawa, 1999) was run within RAxML. The test indicates that the topologies of all six trees are significantly different, suggesting different evolutionary histories for all six viral regions. In the trees based on the early regions (E1–E7), the HR-clade (red) forms a monophyletic clade, separate from the LRclades, whereas in the late region trees, the HR and LR-clades are mixed (Narechania et al., 2005). This suggests that oncogenicity traces along the early region. This is further supported by the observation that the HR and LR-clades are monophyletic (and thus separate) in the oncogene (E6 and E7) trees.

The observation that phylogenetic approaches indicate an association of the early region with oncogenic potential appears to suggest an evolutionary relatedness and origin of viral carcinogenicity. Based on this

Evolution of Human Papillomavirus Carcinogenicity

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hypothesis, the comparison of phylogenetically related types, with distinct epidemiological risks, should allow researchers to tease apart what separates oncogenic HPV types (OTs) from nononcogenic HPV types (NOTs) within the HR-clade.

VI. THE E6 AND E7 VIRAL ONCOGENES As introduced in Section IV, a PV infection requires the host cell machinery to drive its replication. In order to utilize the DNA replication machinery, the virus needs to subvert the cells’ terminal differentiation process. This will result in an increased number of infected cells, and eventually the production of infectious virions. Both the E6 and E7 proteins have been implicated in the stimulation of cell cycle progression. The following is not meant to be an exhaustive listing of E6 and E7 functions, since several recent reviews have been dedicated to both these proteins. We will, however, briefly examine some of these functions (Table I). The ability to degrade p53, thus interfering with the cellular stress response is probably the best-characterized function of the viral E6 protein (Band et al., 1991). HPV16 E6 forms a complex with p53 and the 100 kDa E6AP, an E3 ubiquitin ligase (Scheffner et al., 1993). This interaction results in the ubiquitination of p53 and its subsequent degradation through the proteasomal pathway. However, HPV-induced immortalization of primary keratinocytes does not appear to depend on p53 degradation (Kiyono et al., 1998). However, it is clear that E6-induced degradation of p53 as well as Bak (a proapoptotic protein; Thomas and Banks, 1999) represents a strong antiapoptotic signal. In addition, the increased mitotic activity through E6 interaction with Blk (a SRC family protein kinase) further facilitates acquired mutagenic effects (Oda et al., 1999). Recently, the importance of cell polarity in the development of cancer has started to be appreciated (reviewed in Wodarz and Nathke, 2007). It is likely that the interactions of the E6 protein with cell polarity factors such as the PDZ domain-containing proteins (Thomas et al., 2008) and paxillin (Tong and Howley, 1997; Tong et al., 1997) provide an important mechanism for progression to malignancy. In addition, the degradation of E6TP1, a novel GAP protein, also appears to play an important role in HPV carcinogenesis (Gao et al., 1999). Interestingly, amongst the different functions of E6, it appears that activation of the hTERT promoter (reviewed in Galloway et al., 2005) in combination with the combined perturbation of the p16/pRb pathway is sufficient to immortalize primary HFKs (Kiyono et al., 1998). Nevertheless, additional factors are probably needed for induction of malignancy in infected humans. HPV16 E7 has been shown to preferentially bind to the hypophosphorylated (cell cycle inhibiting) pRb. This results in a decreased

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TABLE I Summary of the interacting partners of the HPV oncoproteins Partner

Function

Reference

E6 Bak

Bcl2 family member

Thomas and Banks (1999) Oda et al. (1999)

Blk E6AP E6TP1 hTERT p53 Paxillin PDZ-containing proteins E7 AP1 c-jun Cullin 2 Cyclin A Cyclin E Histone H1 kinase hTLR9 Mi2B p21 Pocket protein family TAP-1 TBP

src family tyrosin protein kinase E3 Ubiquitin ligase Putative GAP protein Maintains telomere length Tumor suppressor Focal adhesion protein Cellular polarity

Transcription factor Transcription factor E3 Ubiquitin ligase Cell cycle control Cell cycle control Cell cycle control Innate immunity Histone deacetylase complex Cell cycle control Cell cycle control Antigen processing Transcription factor

Rolfe et al. (1995) Gao et al. (1999) Veldman et al. (2001) Werness, et al. (1990) Tong and Howley (1997) Thomas et al. (2008)

Antinore et al. (1996) Antinore et al. (1996) Huh et al. (2007) Arroyo et al. (1993) McIntyre et al. (1996) Davies et al. (1993) Hasan et al. (2007) Brehm et al. (1999) Helt et al. (2002) Helt and Galloway (2003) Vambutas et al. (2001) Massimi, et al. (1997)

interaction of pRb with E2F and progression through the cell cycle (Chellappan et al., 1992). It appears that the complex formed by pRb, E7, and the ubiquitin ligase cullin 2 is responsible for the E7-induced degradation of pRb (Huh et al., 2007). In addition, E7 also interacts with two other members of the pocket protein family, p107 and p130, which also negatively regulate E2F activity (Dyson et al., 1989; Genovese et al., 2008; Helt and Galloway, 2003; Morris et al., 1993). The direct interactions of the E7 protein with the histone H1 kinase (Davies et al., 1993), cyclin A (Arroyo et al., 1993), cyclin E (McIntyre et al., 1996), the histone deacetylase complex Mi2B (Brehm et al., 1999), and several transcription factors (c-jun,

Evolution of Human Papillomavirus Carcinogenicity

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AP1 (Antinore et al., 1996) and TATA binding protein (Massimi et al., 1997)) further allow the virus to abrogate the normal cell cycle control pathways and reenter the DNA synthesis phase. Finally, it has been shown that both pRb and p21 need to be inactivated through E7 expression to circumvent cell cycle arrest (Helt et al., 2002). Interestingly, the expression of E7 stabilizes p21 and p53 levels ( Jones et al., 1997, 1999), so expression of E6 proteins in turn needs to reduce the levels of p53 induced by E7 expression. In addition to modulating cell cycle control, the E7 protein has been implicated in the immune response. The E7 protein downregulates the levels of human toll-like receptor hTLR9 (Hasan et al., 2007) and the ER protein TAP-1 (Vambutas et al., 2001). It was recently shown that variations within the TAP-1 gene were associated with increased risk for highgrade cervical neoplasia (Einstein et al., 2009), suggesting that changing the expression levels of these proteins might play a role in viral immune evasion. Nonetheless, it is important to note that all PVs need to ‘‘kick start’’ the cell cycle, to complete their life cycle. This has to be achieved irrespective of carcinogenic potential. So, it appears unlikely that the tweaking of basic cell cycle regulators is what separates oncogenic (OT) from nononcogenic (NOT) types. We argue that the subtleties of viral oncogenesis are best understood through the comparison of phylogenetically related types with different biological behaviors. We suggest that a thorough understanding of the evolutionary history, as well as the epidemiology of these viruses, should guide the interpretation of biochemical assays.

VII. EVOLUTIONARY HISTORY OF THE E6 AND E7 ONCOPROTEINS The E6 protein is a small basic protein of about 150 amino acids. Every E6 protein contains four CxxC domains, which are involved in the formation of N- and C-terminal zinc-binding domains (Grossman and Laimins, 1989; Fig. 6). The presence of these domains has been shown to be essential for the proteins’ functions. A solution structure of the E6 C-terminal domain has been proposed (Nomine et al., 2003), and was used to suggest that a single-domain protein possessing the same fold might have once existed. It was recently shown that a short protein reminiscent of a single domain E6 (Nomine et al., 2006) was present before reptiles and mammals diverged about 310 MYA (Herbst et al., 2009; Van Doorslaer et al., 2009). The discovery of these short E6 proteins allowed for the experimental confirmation of a hypothesis stipulated over 20 years ago (Cole and Danos, 1987). It appears that the E6 ORF gained access into the viral genome after which an (internal) duplication gave rise to the

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Koenraad Van Doorslaer and Robert D. Burk

A M

CxxC

1

30

M

LxCxE

F CxxC

CxxC RRRETQL

CxxC

Y

HPV16 E6 47

66

A

CxxC

55

66

81

103

139

151 146

CxxC

HPV45 E7 1

26

B

102 106

C -C

C

-

N-

-N

FIGURE 6 E6 and E7 protein structures. (A) The positions of the CxxC-N29-CxxC domains in both proteins are indicated. The CxxC regions highlighted in blue and red correspond to the regions highlighted in (B) and (C), respectively. In E6 amino acid F47 is highlighted, a position implicated in the interaction with E6AP and degradation of p53 (Nomine et al., 2006). Y81 is the most N-terminal residue solved in the structure shown in (B). The region involved in binding to PDZ proteins is indicated in gray and is not shown in the structure in (B). In the E7 model, the pRb binding domain (LxCxE) and the starting point of the structure shown in (C), A55 are indicated. (B) Shows the NMR structure of the C-terminal domain of HPV16 E6 (Nomine et al., 2006). The file was downloaded from the pdb (www.rscb.org/pdb) with accession number 2FK4 and visualized in MacPymol (http://www.pymol.org). Both CxxC regions are highlighted. (C) Shows the NMR structure of the C-terminal domain of HPV45 E7 (Ohlenschlager et al., 2006). The file was downloaded from the pdb (www.rscb.org/pdb) with accession number 2EWL and visualized in MacPymol (http://www.pymol.org). Both CxxC regions are highlighted.

typical E6 protein we observe today in the majority of extant PVs (Van Doorslaer et al., 2009). The absence of an E6 ORF in certain PVs (Chen et al., 2007; Van Doorslaer et al., 2006), suggests a recent loss of this ORF from these viruses. At 100 aa, the E7 protein is shorter than the E6 protein, and it also contains a single CxxC Zn-finger domain (Fig. 6). Sequence analysis of the E6N, E6C, and E7 Zn-binding domains suggests E7 is a duplication of

Evolution of Human Papillomavirus Carcinogenicity

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the ancestral single E6 domain (Van Doorslaer, unpublished) The recently solved E7 structures show a N-terminal region, which includes the conserved pRb-binding motif (consensus LXCXE), followed by an obligate homodimeric 50-residue zinc-binding domain (Liu et al., 2006; Ohlenschlager et al., 2006). Two groups independently showed that the E6 genes mutated at a higher rate compared to the rest of the viral genome (Garcia-Vallve et al., 2005; Rector et al., 2007). Based on the analysis of feline PVs, Rector and colleagues quantified the different mutation rates to be  2.4  10 8 versus 1.95  10 8 nucleotide substitutions/site/year, respectively (Rector et al., 2007). Interestingly, this means that the E6 mutates at basically the same rate as the URR (2.69  10 8). Since the URR is noncoding, one would expect this region to have a high mutational rate and be highly variable (Chen et al., 2005). One of the potential explanations suggested for the elevated E6 mutation rate is that the E6 ORF gained access into the viral genome after the basic genome structure was formed (Garcia-Vallve et al., 2005). The suggested duplication event might also aid in explaining this phenomenon (Van Doorslaer et al., 2009). Alternatively, the E6 ORF might be under less stringent purifying selection due to selective functions necessary for the PV life cycle. The observed incongruencies between the different viral regions (Fig. 5; Garcia-Vallve et al., 2005; Narechania et al., 2005) argue that the PV genome consists of three independently evolving units (E6–E7, E1–E2, and L1–L2), with oncogenicity most closely tracking the evolution of the E6 and E7 ORFs.

VIII. TOWARD A BIOCHEMICAL UNDERSTANDING OF ALPHAPAPILLOMAVIRUS ONCOGENICITY The correlation between the early region phylogeny and viral oncogenicity implies that evolutionary changes conveying carcinogenicity to the most recent common ancestor (MRCA) of the current HR-clade are maintained within today’s extant OT HPVs. As mentioned earlier, previous studies have shown that OT HPV types, HPV16 and HPV18, degraded p53, whereas NOT types HPV6 and -11 were unable to do so (Lechner and Laimins, 1994; Scheffner et al., 1990). Based on these observations, it has long been assumed that p53 degrading ability was strongly associated with the oncogenicity by OT HPVs. In addition, it has been argued that the ability to interact with PDZ domain-containing cellular proteins was a unique feature of OT HPVs (Kiyono et al., 1997; Lee et al., 1997). The importance of degrading PDZ domain-containing proteins for viral oncogenesis was supported by the observation that the E7 protein might complement this function in RhPV1-induced oncogenesis (Tomaic et al., 2009). However, with HPV6

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Koenraad Van Doorslaer and Robert D. Burk

and -11 causing condyloma accuminata and laryngeal papillomas, while HPV16 and -18 primarily infecting the cervix, it is possible that the observed differences in activity are due to tissue tropism differences. This is supported by data showing that skin types associated with nonmelanoma skin cancer do not affect p53 levels (Steger and Pfister, 1992). We need to be careful that the comparison of types that are distantly related does not result in overinterpretation of biochemical data. Figure 7 shows a summary of recent data on how the E6 proteins of different types affect the steady-state levels of p53 and Magi1c and hDLG 58 33 52 35 16 31 73 34 18 45 70 68 39 82 51 26 66 56 53 30 42 40 6 11 44 54 10 90 106 71 62 61 2

Species 9 9 9 9 9 9 11 11 7 7 7 7 7 5 5 5 6 6 6 6 1 8 10 10 10 13 2 15 15 15 3 3 4

Epi. p53 (1) p53 (2) p53 (3) Magi1c (4) hDLG (4) OT Yes Yes No No Yes OT Yes ~ Yes Yes OT Yes No Yes Yes OT Yes Yes Yes Yes OT Yes ~ Yes OT Yes NOT Yes N/A Yes OT Yes No Yes OT Yes ~ Yes Yes NOT Yes No Yes N/A Yes Yes OT Yes Yes Yes N/A Yes No Yes OT ~ Yes Yes Yes NOT Yes Yes NOT No No Yes OT No No Yes Yes NOT Yes Yes N/A No NOT No No No NOT No NOT No No No No NOT No No No NOT No No No NOT No No No No NOT No NOT No NOT No NOT Yes NOT No NOT No No No No NOT No

FIGURE 7 Evolutionary and epidemiologically correlated analysis of p53 and PDZ degradation activities. This figure shows the E6 tree depicted in Fig. 5. Branches were culled when no biochemical data was available. The a-PV species are named according to de Villiers et al. (de Villiers et al., 2004). The epi. column shows the epidemiological classification (oncogenic, OT; nonOT, NOT, insufficient data, N/A) according to Bouvard et al. (2010). The following columns summarize the data presented in p53 (1) (Hiller et al., 2006), p53 (2) (Hiller et al., 2008), p53 (3) (Fu et al., manuscript submitted), Magi1c (4), and hDLG (4) (Muench et al., 2009). In these columns, ‘‘yes’’ indicates degradation of the target, ‘‘no’’ means no degrading potential, and ‘‘’’ signifies intermediate potential. Branches are colored according to Fig. 5, with the red, HR-clade clustering both the OT and NOT types.

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(PDZ-containing proteins; Hiller et al., 2006; Hiller et al., 2008; Muench et al., 2009). This tree, based on the nucleotide sequence of the E6 proteins, clusters all the oncogenic types (Bouvard et al., 2010) in a single (red) HR-clade; however, not all members of this clade actually cause disease. The data clearly show that p53 degradation by OT types is not highly correlated with oncogenicity. On the contrary, it appears that this trait was inherited from the common ancestor of this clade. The most parsimonious explanation would be that the types no longer able to degrade p53 (e.g., HPV40, 6, 11, 44, 54, etc.) lost this ability after the HR MCRA diverged. This suggests that additional functions separate the NOTs from the OTs. Based on Fig. 7, the PDZ-proteins might be interesting candidates. Degradation of Magi1c appears to recapitulate a significant portion of the epidemiological classification of OT HPVs. However, the use of the mouse homolog of Magi1c (Dobrosotskaya et al., 1997) in an in vitro assay (Muench et al., 2009) might obscure the interpretation of the data, especially when compared to the human hDLG used in the same study. However, the data do warrant a detailed investigation into the effects of phylogentically related viral types on PDZ-containing proteins. The goal of these comparative genomic studies should be to identify the biochemical functions that separate the OT from the NOTs. Since both E6 and E7 are necessary to successfully transform primary keratinocytes (Munger et al., 1989), it is possible that the OT-defining functionality lies within the E7 protein and/or an interaction between the E6 and E7 proteins, introducing a level of complexity into the biological system. Of interest is the observation that two natural variants of HPV16 E6 (from the European prototype (E(p) and Asian American (AA) (nonEuropean) lineages (Chen et al., 2005)) affect in vitro transformation differently in a manner corresponding to their differences in epidemiological risk classification (Richard et al., 2010). The authors used prototype E7 in conjunction with E(p) E6 or AA E6 (Q14H/H78Y/L83V) in an array of functional studies. They observed no differences in p53 inactivation or telomerase activation. However, only the AA variant was able to successfully transform the primary HFKs. In addition, proteomic analysis showed markedly different signatures between both sets of cell lines (Richard et al., 2010). Not only did this study suggest an important role for the E6 protein, but it also illustrated beautifully how an understanding of the epidemiology and evolutionary path of these viruses can guide in vitro experiments.

IX. CONCLUSIONS AND PERSPECTIVES In the present review, we argue that the identification of functions associated with HPV oncogenic potential should be done carefully and with a consideration of viral epidemiology and evolutionary history.

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More detailed investigations have suggested that the differences between OT and NOT a-papillomavirus types are more complex than originally anticipated. Phylogenetic analysis emerged from the field of systematics, where it added an extra dimension to anatomical trait analysis for species classification. However, phylogeny has gained access into all fields of biology, having been applied to understand virulence (Hon et al., 2006), drug resistance (Seret et al., 2009), the origin of epidemics (Smith et al., 2009), and forensic investigations (Arens, 1999). We argue that the wealth of HPV sequence and epidemiological data now opens the door for HPV VWAS (viral genome wide association studies). The goal of human GWAS studies is to identify SNPs associated with a specific phenotype or disease. The 8 kb size of the HPV viral genome, coupled with excellent epidemiological data and the extreme oncogenic risk of some HPV types but not others, make HPV genome analyses an excellent model system to dissect the genetic contribution of viral nucleotides to disease. We also point out that the extant human genital HPVs show a strong correlation between the epidemiological classification and the evolutionary history of their oncogenic proteins. Building on these arguments, we believe that evolutionary analyses will allow researchers to identify those evolutionary changes potentially conferring carcinogenic potential to the OTs. These identified SNPs can next be manipulated in vitro and in vivo to qualitatively test the differences between these proteins. Although an effective HPV vaccine for HPV types 6, 11, 16, and 18 is currently available, the world population of women will be at risk for cervix cancer for generations to come. This is due to the high cost of the vaccine, the lack of therapeutic activity in women currently infected, and the low penetration of vaccine in the most at-risk population in the developing world. Moreover, cervix cancer is a solid tumor that has a complex biology, given the average 20–40 years of development from the time of infection. Further studies are warranted not only for the public health benefits, but also for the potential to investigate the mechanisms of HPV pathogenicity and viral behavior.

ACKNOWLEDGMENTS The authors gratefully acknowledge past and current members of the Burk lab for their contributions to the research efforts of the group, in particular, Dr Zigui Chen, for the preparation of Fig. 1. We are also grateful to the contributions of all our colleagues in the PV field and apologize that we could not include all recent discoveries in this review. This work was supported in part by Public Health Service awards CA78527 from the National Cancer Institute (R. D. B.) and center grants to the Einstein Cancer Research Center and the Center for AIDS Research (CFAR).

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Tong, X., Salgia, R., Li, J. L., Griffin, J. D., and Howley, P. M. (1997). The bovine papillomavirus E6 protein binds to the LD motif repeats of paxillin and blocks its interaction with vinculin and the focal adhesion kinase. J. Biol. Chem. 272(52):33373–33376. Vambutas, A., DeVoti, J., Pinn, W., Steinberg, B. M., and Bonagura, V. R. (2001). Interaction of human papillomavirus type 11 E7 protein with TAP-1 results in the reduction of ATPdependent peptide transport. Clin. Immunol. 101(1):94–99. Van Doorslaer, K., Rector, A., Vos, P., and Van Ranst, M. (2006). Genetic characterization of the Capra hircus papillomavirus: A novel close-to-root artiodactyl papillomavirus. Virus Res. 118(1-2):164–169. Van Doorslaer, K., Sidi, A. O., Zanier, K., Rybin, V., Deryckere, F., Rector, A., Burk, R. D., Lienau, E. K., van Ranst, M., and Trave, G. (2009). Identification of unusual E6 and E7 proteins within avian papillomaviruses: Cellular localization, biophysical characterization, and phylogenetic analysis. J. Virol. 83(17):8759–8770. Veldman, T., Horikawa, I., Barrett, J. C., and Schlegel, R. (2001). Transcriptional activation of the telomerase hTERT gene by human papillomavirus type 16 E6 oncoprotein. J. Virol. 75(9):4467–4472. Watts, S. L., Ostrow, R. S., Phelps, W. C., Prince, J. T., and Faras, A. J. (1983). Free cottontail rabbit papillomavirus DNA persists in warts and carcinomas of infected rabbits and in cells in culture transformed with virus or viral DNA. Virology 125(1):127–138. Werness, B. A., Levine, A. J., and Howley, P. M. (1990). Association of human papillomavirus types 16 and 18 E6 proteins with p53. Science. 248(4951):76–79. Wodarz, A., and Nathke, I. (2007). Cell polarity in development and cancer. Nat. Cell Biol. 9(9):1016–1024. Yang, L., Li, R., Mohr, I. J., Clark, R., and Botchan, M. R. (1991). Activation of BPV-1 replication in vitro by the transcription factor E2. Nature 353(6345):628–632. Zhao, K. N., Sun, X. Y., Frazer, I. H., and Zhou, J. (1998). DNA packaging by L1 and L2 capsid proteins of bovine papillomavirus type 1. Virology 243(2):482–491. Zhao, K. N., Hengst, K., Liu, W. J., Liu, Y. H., Liu, X. S., McMillan, N. A., and Frazer, I. H. (2000). BPV1 E2 protein enhances packaging of full-length plasmid DNA in BPV1 pseudovirions. Virology 272(2):382–393. Zhou, J., Liu, W. J., Peng, S. W., Sun, X. Y., and Frazer, I. (1999). Papillomavirus capsid protein expression level depends on the match between codon usage and tRNA availability. J. Virol. 73(6):4972–4982.

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3 Influenza Vaccines: The Good, the Bad, and the Eggs Stacey Schultz-Cherry and Jeremy C. Jones

Contents

Abstract

I. II. III. IV. V. VI.

Influenza: The Burden The Virus and Its Tricks The Good, the Bad, and the Eggs Pandemic Vaccine: Success in 2009 On the Horizon: Bird Viruses The Future of Influenza Vaccines: Teaching an Old Dog New Tricks VII. The Quest for a ‘‘Universal’’ Influenza Vaccine: Is it Possible? VIII. Conclusions References

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Outbreaks of influenza A viruses continue to cause morbidity and mortality worldwide. The global disease burden of influenza is substantial. While antiviral therapies are available, influenza vaccines are the mainstay of efforts to reduce the substantial health burden from seasonal influenza. Inactivated influenza vaccines have been available since the 1940s, with live attenuated, cold-adapted vaccines becoming available in the United States in 2003. In spite of the successes, more research is needed to develop more effective seasonal influenza vaccines that provide long-lasting immunity and broad protection against strains that differ antigenically from vaccine viruses. This review introduces the virus and its disease, the current state of seasonal and pandemic influenza vaccines, and the challenges we face in the future.

Department of Infectious Disease, St. Jude Children’s Research Hospital, Memphis, Tennessee, USA Advances in Virus Research, Volume 77 ISSN 0065-3527, DOI: 10.1016/S0065-3527(10)77003-2

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2010 Elsevier Inc. All rights reserved.

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I. INFLUENZA: THE BURDEN Influenza A is a highly communicable respiratory disease transmitted primarily via respiratory droplets that are expelled by coughing or sneezing of infected individuals. It can also be spread through contact with surfaces containing droplets from infected individuals, followed by touching one’s mouth or nose (CDC, 2005). In the United States alone, 5–20% of the population acquires influenza on an annual basis, resulting in approximately 200,000 hospitalizations and 36,000 deaths (Barker and Mullooly, 1980; Thompson et al., 2004; Tosh et al., 2010). Much of the morbidity and mortality of seasonal influenza results from complications of infection rather than from primary viral pneumonia (Rothberg and Haessler, 2010). Influenza infections are the cause of yearly epidemics as the virus spreads through populations, which primarily occur in the Northern Hemisphere from late December through early March (CDC, 2005). It has been proposed that temperature and humidity influence transmission of influenza, and may explain seasonal spread as opposed to infections occurring year-round (Lowen et al., 2007; Schaffer et al., 1976). In addition to seasonal epidemics, influenza A viruses cause pandemics. There were at least three documented influenza pandemics in the twentieth century; the most notorious being the 1918 Spanish Flu, which caused 40–50 million deaths worldwide (Nguyen-Van-Tam and Hampson, 2003; Taubenberger et al., 2001; Tumpey et al., 2005; Yewdell and Garcia-Sastre, 2002). Beginning in April 2009, we faced our first influenza pandemic of the twenty-first century from a novel swine-origin H1N1 virus (Peiris et al., 2009a,b; Tang et al., 2010). Clinically, the signs and symptoms of 2009 H1N1 influenza are similar to those of seasonal influenza; fever, head and muscle ache, malaise, nonproductive cough, and sore throat that manifest 24–96 h postinfection. The incubation period ranges from 2 to 7 days with the infectious period, characterized by viral shedding, ranging from 1 day before symptom onset to at least 5–7 days after onset (Novel Swine-Origin Influenza A virus Investigation Team, 2009). Unlike seasonal influenza, 25–40% of people infected with the 2009 H1N1 pandemic virus developed diarrhea and vomiting ( Jain et al., 2009; Novel Swine-Origin Influenza A Investigation Team, 2009; Swedish et al., 2010). The majority of 2009 H1N1 influenza infections have been selflimiting and mild in nature, although severe illness has been reported (Napolitano et al., 2010). For the first time, obesity has emerged as a risk factor for developing severe 2009 H1N1 influenza infection (Rothberg and Haessler, 2010).

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II. THE VIRUS AND ITS TRICKS Influenza is an enveloped, single-strand, negative-sense RNA virus of the family Orthomyxoviridae (Lamb and Krug, 2001). The Orthomyxoviruses are subdivided into five genera, influenza A, B, and C, Thogotoviruses, and salmon anemia virus (Lamb and Krug, 2001). Influenza A viruses infect a wide range of animals, including humans, birds, pigs, horses, cats, and many others, and are associated with seasonal epidemics and pandemics. Influenza B infects a smaller number of species and is a substantial cause of annual epidemics. The C viruses rarely account for human infections and epidemics. Thus, the review will focus on influenza A viruses. The genome of influenza A virus is composed of eight negative-sense RNA segments which encode 10 to 11 proteins (Fig. 1; Conenello and Palese, 2007; Lamb and Krug, 2001; Shaw et al., 1992; Yewdell and GarciaSastre, 2002). The virus is 80–120 nm in size, with individual virions ranging in shape from spherical to pleomorphic (Lamb and Krug, 2001; Nicholson et al., 2003). Two viral-encoded glycoproteins are located within the envelope: hemagglutinin (HA) and neuraminidase (NA). HA binds the virus receptor, while NA enzymatically cleaves receptors to

Negativesense ssRNA M2 PA

PB1

PB2

M1 NA NP

PB2 PB1 PA HA NP NA M NS

HA

FIGURE 1 Influenza Virion. Used with permission from Nature Publishing Group. From Horimoto and Kawaoka (2005). Each segment has only one copy of PB2, PB1, and PA.

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mediate virus exit (Lamb and Krug, 2001). Influenza A viruses are subtyped based on the major subtype of HA and NA expressed on the virion (e.g., H3N2, H5N1). To date, 16 antigenically distinct HA proteins and nine distinct NA proteins have been identified (Horimoto and Kawaoka, 2005; Renegar, 1992). The HA protein binds to sialic acid moieties linked to an underlying galactose (Sawada et al., 2007; Skehel and Wiley, 2000). The linkage of the sialic acid to the underlying carbohydrate is thought to influence the host range of influenza viruses. a2–6 linkages are prevalent on epithelial cells of the upper respiratory tracts of humans and other mammals, while a2–3 linkages are more common on host cells of the gastrointestinal tract of avian species (Suzuki, 2005). This dichotomy has proved less acceptable with the discovery of a2–3 linkages in the human lung and a2–6 linkages in tissue from chickens and quail. However, influenza viruses are often still characterized as mammalian or avian, and a loosely defined ‘‘species barrier’’ exists due to the decreased prevalence of a receptor linkage in a particular host (Ito and Kawaoka, 2000). A third membrane protein, M2, serves as an ion channel and functions during virus uncoating (Pinto and Lamb, 2006; Wang et al., 1993). Underlying the envelope is a layer of matrix protein (M1) that provides structural integrity (Gregoriades and Frangione, 1981; Nayak et al., 2004). Each genomic segment is coated with several copies of nucleoprotein (NP) and one copy of each of the three polymerase genes, PA, PB1, and PB2 (Lamb and Krug, 2001; Shaw et al., 1992). Collectively, this structure is known as the ribonucleoprotein complex (RNP) (McCauley and Mahy, 1983). The nuclear export protein (NEP), previously referred to as NS2, interacts with RNPs to facilitate packaging late in infection (Lamb and Krug, 2001). The nonstructural protein expressed during infection, NS1, plays a role in immune evasion (Krug et al., 2003; Lipatov et al., 2005; Talon et al., 2000; Wang et al., 2000). The recently identified PB1-F2 protein is encoded from an alternative reading frame in the PB1 polymerase gene and functions as a virulence factor through an unidentified mechanism of activity (McAuley et al., 2007, 2010). Circulating influenza viruses are constantly mutating. Minor antigenic changes due to the error-prone nature of the virally encoded RNA-dependent RNA polymerase is called antigenic drift (Fig. 2). The changes that occur to HA and NA help the virus evade the immune response in a host population whether through prior infection or vaccination and are the basis for annual influenza epidemics. For this reason, influenza vaccines are frequently reformulated to account for viral evolution. Although antigenic drift is problematic, antigenic shift is of greater concern. The influenza virus genome is segmented, allowing for the reassortment of gene segments in dually infected cells (Fig. 3). Given the widespread tropism of influenza A viruses and the potential for zoonotic transmission, coinfection with at least two different influenza A viruses can occur,

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1 Each year’s flu vaccine contains three flu strainstwo A strains and one B strain-that can change from year to year. 2 After vaccination, your body produces infection-fighting antibodies against the three flu strains in the vaccine. Antibody

3 If you are exposed to any of the three flu strains during the flu season, the antibodies will latch onto the virus’s HA antigens, preventing the flu virus from attaching to healthy cells and infecting them. 4 Influenza virus genes, made of RNA, are more prone to mutations than genes made of DNA. Viral RNA

Mutation

Antibody

Link studio for NIAID

HA antigen

5 If the HA gene changes, so can the antigen that it encodes, causing it to change shape. HA gene HA antigen

Antibodies

6 If the HA antigen changes shape, antibodies that normally would match up to it no longer can, allowing the newly mutated virus to infect the body’s cells. This type of genetic mutation is called “Antigenic drift.”

FIGURE 2 Influenza Antigenic Drift. Image courtesy of the National Institute of Allergy and Infectious Diseases (NIAID) http://www.niaid.nih.gov/topics/Flu/Research/basic/ Pages/AntigenicDriftIllustration.aspx.

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The genetic change that enables a flu strain to jump from one animal species to another, including humans, is called “Antigenic shift.” Antigenic shift can happen in three ways:

Bird host

B Without undergoing Bird influenza A strain genetic change, a bird strain of influenza A can jump directly from a duck or other aquatic bird to humans.

HA antigen

C Without undergonig genetic change, a bird strain of influenza A can jump directly from a duck or other aquatic bird to an intermediate animal host and then to humans.

NA antigen

The new strain may further evolve to spread from person to person. If so, a flu pandemic could arise.

Human host

Human influenza A strain

A-1 A duck or other aquatic bird passes a bird strain of influenza A to an intermediate host such as a chicken or pig.

A-2 A person passes a HA NA human strain of antigen antigen influenza A to the same chicken or pig. (note that reassortment can occur in a person who is infected with two flu strains.) A-3 When the viruses infect the same cell, the genes from the bird strain mix with genes from the human strain to yeild a new strain.

Viral entry intermediate host cell

A-4

New influenza strain

The new strain can spread from the intermediate host to humans.

Intermediate host cell Genetic mixing Link studio for NIAID

Intermediate host (pig)

FIGURE 3 Influenza Antigenic Shift. Image courtesy of the National Institute of Allergy and Infectious Diseases (NIAID) http://www.niaid.nih.gov/topics/Flu/Research/basic/ Pages/AntigenicShiftIllustration.asp.

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as evidenced by the 2009 H1N1 pandemic virus (Garten et al., 2009). This is a triple-reassortant virus containing segments from avian, swine, and human lineages that may have undergone a reassortment event 10–17 years ago (Ilyushina et al., 2010; Smith et al., 2009a,b). Although only three subtypes of influenza A virus are known to efficiently transmit in humans (H1, H2, and H3), viruses representing all 16 HA subtypes are found in aquatic birds (Krauss et al., 2004), which serve as the major reservoir. Reassortment between any of these avian strains and the right combination of mammalian segments could result in a new pandemic virus. Thus, we must remain vigilant in surveillance efforts to identify new influenza viruses in nature.

III. THE GOOD, THE BAD, AND THE EGGS Though antiviral drugs exist to treat influenza infection, primary control of yearly epidemics is mediated through comprehensive vaccination programs (Cox et al., 2004; Stephenson and Nicholson, 2001). Currently, two forms of influenza vaccine are available for widespread distribution: inactivated vaccines (INV) and live-attenuated vaccines (LAV). INV consist of virus of a chosen subtype propagated in embryonated chicken eggs, purified, and detergent split (Girard et al., 2005). Multiple subtypes of influenza A (H3N2 and H1N1) as well as influenza B are mixed to provide a broader range of protection against circulating viruses, and these vaccines are administered intramuscularly on a yearly basis (Stephenson and Nicholson, 2001). INV produces both systemic and localized antibody responses, but the predominant response is serum neutralizing IgG against the surface HA and NA proteins. Production of neutralizing serum IgG is a commonly accepted correlate to protection against influenza infection, and it is believed to inhibit disease progression and severity. However, INV are less effective in eliciting production of secretory IgA and often do not provide heterosubtypic immunity (protection against one or more nonrelated subtypes; Cox et al., 2004; Lu et al., 2006a,b). INV are approved for children, the elderly, and those with certain chronic medical conditions, but must be reformulated and redelivered on an annual basis due to genetic variance in the virus genome as it circulates through populations (CDC, 2007). LAV contains the same subtype viruses as the inactivated vaccine. However, the viruses are cold adapted, inhibiting replication at temperatures exceeding 25  C, thus restraining replication to the upper airway (McCarthy and Kockler, 2004). LAV are delivered intranasally, inducing a mucosal IgA response that is believed to limit early viral replication, prevent further disease progression, and may afford heterosubtypic immunity (Cox et al., 2004; Nichol and Treanor, 2006; Stephenson and

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Nicholson, 2001). Since it is composed of live virus, vaccine safety is guaranteed for a reduced portion of the population (ages 5–49). High-risk groups, such as pregnant women, the elderly, and immunocompromised, are also excluded (CDC, 2007). Additionally, viral shedding, rhinorrhea, sore throat as well as other flu-like symptoms are typical side effects (McCarthy and Kockler, 2004). The process for generating both inactivated and live-attenuated vaccine for seasonal influenza is well defined. Twice a year, representatives from the four collaborating World Health Organization (WHO, 2006) Centers for Reference and Research on Influenza and key national laboratories meet to review surveillance data and decide on the appropriate strains for inclusion in the following season’s influenza vaccine (WHO, 2009a). Two influenza A strains (one H1N1 and one H3N2) and an influenza B strain are recommended. Strain selection may differ in the northern and Southern hemispheres depending on the currently circulating viruses. Several factors are considered when making recommendations, including (1) predicting which strains are likely to cause the following season’s epidemic based on surveillance information, (2) the antigenic similarity of the chosen vaccine strain to the predicted circulating strain, (3) the immunogenicity of the selected strain, and (4) the suitability of the strain for vaccine production (i.e., does it grow to high titers in eggs?). The entire process from prediction to production/delivery takes 6–8 months. There are several pitfalls that could lead to decreased vaccine efficacy or even shortages. If the vaccine component is not well-matched to the circulating strain, which could be undergoing antigenic drift during the process, efficacy will be decreased. This occurred during the 2007–2008 influenza season when an A/Wisconsin/67/2005 (H3N2)-like virus was recommended for the vaccine, while an A/Brisbane (H3N2)-like virus became the predominant circulating strain (CDC, 2007–2008). Supply shortages have resulted from contamination of embyronated eggs, reduction in the numbers of vaccine manufacturers, and earlier than expected outbreaks. Vaccine production is also highly dependent on an adequate supply of eggs. On average, approximately one egg is required to produce one dose of one vaccine strain. In spite of the pitfalls, required guess work, and long production time, the majority of the people in the United States have access to seasonal influenza vaccines (Tosh et al., 2010).

IV. PANDEMIC VACCINE: SUCCESS IN 2009 Experts questioned whether a new vaccine could be developed and administered with sufficient speed during a pandemic. Our experience with the 2009 H1N1 pandemic suggests that the answer is yes, but success may be dependent on the virus and country (Oshitani et al., 2008).

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Since the emergence of the highly pathogenic H5N1 influenza viruses in humans, a great deal of research and resources has gone into pandemic preparedness and vaccine development (Azziz-Baumgartner et al., 2009). This groundwork may have been invaluable in dealing with the first influenza pandemic of the twenty-first century, due not to the predicted H5N1 avian viruses, but instead to a triple-reassortant virus with swine origins. As early as March 2009, cases of respiratory illness were documented in Mexico (Lopez-Cervantes et al., 2009). Due to the increasing numbers of people displaying influenza-like symptoms, the General Directorate of Epidemiology (DGE)-notified hospitals and influenza surveillance units began documenting and collecting samples from symptomatic patients. The first confirmed case (RT-PCR positive) of influenza A virus was reported on April 23 (Lopez-Cervantes et al., 2009). The number of cases in Mexico continued to increase through the end of April, with a total number of 1918 suspected influenza A infections and 84 deaths (CDC, 2009a). During this time, the Centers for Disease Control and Prevention (CDC) confirmed cases of a novel H1N1 virus in two children in California (CDC, 2009b). Genetic analysis of the Mexican and US viruses confirmed that they were genetically distinct from seasonal H1N1 influenza and contained gene segments from swine, avian, and human origins (Shinde et al., 2009). The number of cases quickly increased throughout Mexico and the United States, leading to both governments declaring public health emergencies. By May 2009, the virus had spread globally, and on June 11, 2009, the WHO officially declared a pandemic (Chan, 2009). By July 2009, the virus had rapidly disseminated worldwide, and it was evident that containment was not a feasible option. Strategies focused on monitoring virus evolution and the production of a safe and effective vaccine. Early serological data demonstrated that the immunity provided by seasonal influenza vaccine would be ineffective against the current pandemic virus (CDC, 2009c; Hancock et al., 2009). Although recent studies suggest that cytotoxic T cells established by seasonal influenza may cross-react against the 2009 H1N1 virus, (Tu et al., 2010) and people that received the 1976 "Swine Flu" vaccine or prior infection have enhanced neutralization responses to the 2009 pandemic virus (Kash et al., 2010; McCullers et al., 2010). Regardless, early data suggested that a new vaccine had to be rapidly produced. Creating a vaccine in a matter of months was a daunting task (Butler, 2009; Collin and de Radigue`s, 2009). Due to diligent surveillance efforts, the WHO made recommendations for vaccine seed strains in late May 2009. During this time, there was little evidence of antigenic drift (Glinsky, 2010). Had the virus proved as mutable as the highly pathogenic H5N1 viruses, recommendation of seed stocks a mere 2 months after virus emergence may have been difficult. However, the virus replicated poorly in embryonated chicken eggs, with

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yields as low as 25% compared to traditional seasonal influenza vaccine, creating a setback in vaccine production (Novartis, 2009a). Faced with potential supply shortages and high public demand, the WHO and CDC formulated a priority-based vaccine campaign. In this scheme, health care workers and high-risk groups such as pregnant women, the elderly, and young adults would be vaccinated first (CDC, 2009d; WHO, 2009b). As production and availability of vaccine improved, inoculation of those outside the defined groups would begin. Despite initial setbacks in production, generation of higher yielding seed viruses allowed production of pandemic H1N1 vaccine to continue throughout the summer of 2009 (Novartis, 2009a). The United States Food and Drug Administration approved Sanofi-Pasteur, CLS Limited, Novartis, and MedImmune to begin production of the monovalent pandemic vaccine, which would be available in both inactivated and live attenuated form. Clinical trials conducted through late summer and early fall demonstrated that the one dose of the pandemic vaccine was immunogenic in the general population. A booster vaccination would be required for children 9 years of age (Greenberg et al., 2009; Kelly and Barr, 2010; Plennevaux et al., 2010; Vajo et al., 2010; WHO, 2009c). By the end of 2009, the fears of supply shortages had abated, and the CDC recommended vaccination for all individuals. Assessment of the outcomes of the pandemic and the degree that vaccination decreased spread and overall morbidity is difficult due to underreporting (Lipsitch et al., 2009; Swedish et al., 2010; WHO, 2010a). As of April 2010, the CDC estimates that 43–88 million people in the United States were infected with the 2009 H1N1 virus, resulting in  8700 to  18,000 deaths. In March 2010, it was announced that the A/California/7/2009 virus would be the H1N1 component of the seasonal vaccine (Anonymous, 2010a). Thus, the production of an immunogenic vaccine in the face of a pandemic was successful. This process was likely expedited by coordination between public health entities, vaccine manufacturers, influenza experts, and local healthcare providers on a global scale. However, would we have been successful if the pandemic had been caused by the highly pathogenic H5N1 viruses?

V. ON THE HORIZON: BIRD VIRUSES Under constant surveillance are the highly pathogenic avian (HPAI) H5N1 influenza viruses that first crossed the species barrier in 1997, causing severe morbidity and mortality in infected humans (Belshe, 1998; Claas et al., 1998; Katz et al., 2000; Lipatov et al., 2004; Peiris et al., 2004; Shortridge et al., 1998; Snacken et al., 1999; Tumpey et al., 2000). Since 1997, the HPAI H5N1 viruses have dramatically increased their host range in both birds and humans, with the viruses becoming endemic in

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domestic poultry in many parts of the world. As of April 2010, contact with infected poultry has led to 495 human infections, resulting in 292 deaths: a mortality rate of 59% (WHO, 2010a,b). The virulence associated with mammalian infections has been linked to exacerbated cytokine responses (Chan et al., 2005; Cheung et al., 2002; Chotpitayasunondh et al., 2005; Lipatov et al., 2005; Szretter et al., 2007) and enhanced virus replication (Shinya et al., 2004), leading to considerable lung damage (Anonymous, 2010b,c; Neumann et al., 2010; Peiris, 2009; Peiris et al., 2009a,b; Uyeki, 2009). Given the significant morbidity, increasing host range and spread, and lack of prior immunity, much attention has been paid to developing treatments and control techniques to limit HPAI H5N1 spread should these viruses emerge on a global scale (Anonymous, 2010d; Keitel and Atmar, 2009). In contrast to the 2009 H1N1 virus, there are several challenges to developing vaccines against the HPAI H5N1 viruses. First, H5N1 influenza viruses are continually evolving, adding to the difficulty of predicting the exact strain that will cause a pandemic (Guan et al., 2009; Li et al., 2004; Lu et al., 2006a,b; Smith et al., 2009a,b; Yen and Webster, 2009). Second, currently circulating HPAI H5N1 viruses are associated with severe disease, increasing the urgency for vaccine administration. Third, the H5N1 are highly pathogenic to chicken embryos, compromising the propagation of high-titer viral stocks (Ehrlich et al., 2008; Wright, 2008). Additionally, these viruses are lethal to domestic poultry, further reducing the availability of large numbers of eggs for vaccine production. Finally, initial studies in mice, ferrets, and human trials indicate that the split-virion vaccines, while effective, may require higher doses of antigen and multiple inoculations to initiate protective immunity (Leroux-Roels et al., 2007; Lin et al., 2006; Lu et al. 2006a,b; Nichol and Treanor, 2006). Regardless, numerous H5N1 vaccine viruses have been proposed and developed based on the available antigenic and epidemiologic data (WHO, 2010b). Five egg-derived, inactivated prepandemic H5N1 influenza vaccines have already received marketing authorization from different regulatory agencies worldwide. Four of them are whole-virion vaccines containing between 6 and 15 mg of hemagglutinin (HA) antigen and adjuvanted with aluminium, while the fifth vaccine is split-virion formulation containing 3.8 mg of antigen with ASO3, a novel oil-in-water emulsion-based adjuvant. A recent study by Prieto-Lara and Mendez (2010) provides a systemic review of the safety and immunogenicity of the H5N1 influenza vaccines licensed to date. Given the constant evolution of the H5N1 viruses and the lack of protection against distinct clades, it is possible that additional H5N1 candidate vaccine viruses may need to be developed. Also of concern are the low pathogenic avian H9N2 viruses. These viruses are also evolving and fall within a number of genetically defined HA lineages, the majority belonging to the G1 and Y280 clades (WHO,

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2010b). Phylogenetic analyses suggest that the H9N2 viruses may have contributed to the generation of the HPAI H5N1 viruses. The H9N2 clade G1 viruses share six viral genes (PB2, PB1, PA, NP, M, and NS) with the 1997 H5N1 viruses associated with human disease (Guan et al., 1999). These viruses are widespread in poultry across Asia and Europe and have repeatedly infected humans, albeit causing a mild disease (Butt et al., 2005; Lin et al., 2000; Peiris et al., 1999). The H9N2 viruses also caused infection in pigs and induced severe disease in experimentally infected mice without prior adaptation (Deng et al., 2010; Hossain et al., 2008). Several recent studies demonstrated that the H9N2 viruses replicate and induce specific responses in human lung cells (Lee et al., 2010; Xing et al., 2010). As past pandemics were not caused by highly pathogenic avian influenza viruses, the endemic of H9N2 viruses in poultry as well as their tropism for humans are at least as likely to cause the potential pandemic as the H5N1 virus. Furthermore, an H9N2 avian–human reassortant virus has been shown to have enhanced replication and efficient transmission in ferrets (Wan et al., 2008). Thus, H9N2 virus group is regarded by the World Health Organization as a potential pandemic candidate (WHO, 2010b). Currently, there are several available vaccine viruses, with the A/Hong Kong/33982/2009 clade G1 virus being the proposed vaccine virus (WHO, 2010b) with the possibility that more candidate viruses may have to be developed in the future. However, the H9N2 viruses are not highly pathogenic to chicken embryos or lethal to domestic poultry, so production in eggs should not be problematic. Several studies have demonstrated that both the INV and LAV H9N2 vaccines were well tolerated and immunogenic in humans, inducing good serum neutralizing antibody responses after receiving two doses of vaccine (Atmar et al., 2006; Hehme et al., 2002; Karron et al., 2009; Stephenson et al., 2003). Unexpectedly, both the Karron and Atmar studies found evidence for H9 seroprevalence in people born after 1968–1970 (Atmar et al., 2006; Karron et al., 2009). These data suggest that recipients of experimental H9N2 influenza vaccines should be screened for the presence of preexisting H9 HI antibody. Despite the success, improvements in production, immunogenicity, and heterosubtypic immune responses are warranted to compete with this ever-changing virus.

VI. THE FUTURE OF INFLUENZA VACCINES: TEACHING AN OLD DOG NEW TRICKS Though the influenza vaccination campaigns have significantly impacted and improved public health, they are not without drawbacks. As discussed previously, several factors contribute to the limited efficacy of current influenza vaccines, including virus mutability, failure to induce

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long-term protection, production hurdles, and decreased immunogenicity in high-risk groups. Thus, the development of improved influenza vaccines that address these key limitations continues. Though a multitude of approaches are currently being considered, it may be a combination of several that is ultimately needed to develop an influenza vaccine that provides long-term, widespread protection against a virus that will likely persist in the human population for the foreseeable future. Propagation in embryonated chicken eggs remains the dominate technique for large-scale production of influenza virus vaccines and require dedicated facilities. The process has provided a safe and effective product for decades. The downfalls of the procedure, as discussed previously, suggest the need to move beyond the egg-based manufacturing system. Cell-based cultures have several advantages over eggs (Brands et al., 1999; Kistner et al., 1999; Minor et al., 2009). The facilities could be used for other purposes, and production is scalable. Validated cell banks can be controlled and approved cell lines readily stockpiled and tracked to standardize the manufacturing process, eliminating the irreproducibility that is often seen with varying lots of chicken eggs. It will also allow vaccination of persons with egg allergies; one of the most frequent food allergies in children under the age of three (Benhamou et al., 2010). Certain viruses may grow better in cells than in eggs. Thus, high-growth reassortant viruses may not be needed and production could begin much sooner, leading to a vaccine that is better matched to the circulating virus. Generation of successful cell-culture-based influenza vaccines using whole virus and virus-like particles has advanced rapidly throughout the past decade (WHO; Doroshenko and Halperin, 2009; Genzel and Reichl, 2009; WHO, 2007). Novartis is currently licensed for production and distribution of their cell-derived subunit influenza vaccine in the European Union (WHO; Novartis, 2009b). Recently, they announced construction of a US-based plant dedicated solely to production of cell-culture-generated vaccines (Novartis, 2009c). Several of the other vaccine manufacturers are in various stages of clinical trials and application for cell-culture-based vaccines in the United States. However, there are several potential issues with cell-culture-based methods. First, cells may be less selective for growth of all strains of influenza virus, which may be problematic for both surveillance and production. Further research is required to determine the optimal cell(s) for influenza propagation using a variety of strains. Second, the production strains should be free of significant extraneous agents, and the cells used in isolation may be contaminated or grow contaminating viruses in the sample in a way that eggs do not. Third, influenza vaccine potency is determined by comparison with a homologous reference preparation, and the egg-grown materials may differ. Fourth, the cell lines developed for the manufacturing process may not be readily licensed to independent

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companies for use due to intellectual property issues. Finally, the WHO estimates that cell-based vaccines may be more expensive to produce, a factor that may be overcome with increasing number of facilities and improvement upon the existing process as cell-based vaccines become mainstream (WHO, 2007; WHO). Another approach to improve vaccine efficacy, especially with poorly immunogenic viruses, is the use of adjuvants. Adjuvants induce stronger protective immune responses and/or lower the dose of antigen required for protection, leading to antigen sparing (Ellebedy and Webby, 2009; Nichol and Treanor, 2006; Tritto et al., 2009). They work through several mechanisms, including concentrating the antigen at the site of injection, leading to prolonged exposure to antigen-presenting cells and by enhancing local responses, potentially by stimulating the inflammasome (Mosca et al., 2008; Nichol and Treanor, 2006; Spreafico et al., 2010). Although not used in the United States (Baylor and Houn, 2009), adjuvants are included in influenza vaccines in many parts of the world. The primary adjuvants in use are the oil-in-water emulsions MF59 (Novartis) and AS03 (GlaxoSmith Kline). Both have shown considerable dose-sparing properties with pandemic H1N1 and H5N1 viruses (Atmar and Keitel, 2009; Banzhoff et al., 2008; Dormitzer et al., 2009; Keitel et al., 2010) and were included in the 2009 monovalent H1N1 vaccine in the European Union (Keitel and Atmar, 2009). Other types of adjuvants currently under investigation for influenza vaccines include toll receptor agonists (CpG DNA, Poly IC double-stranded RNA, flagellin; Cooper et al., 2004; Ichinohe et al., 2009; Skountzou et al., 2010; Wong et al., 2009; Wu et al., 2009), lipid-based candidates (ISCOMs and lipid A; Baldridge et al., 2000; Baldwin et al., 2009; Keitel and Atmar, 2009), and antimicrobial peptides (Fritz et al., 2004). The use of adjuvants in the United States represents a plausible option for improving our current influenza vaccines and may become necessary in the event of another pandemic, especially if the virulence exceeds that of the 2009 H1N1 pandemic. Though, strict regulatory and safety issues will need to be addressed, much can be learned from the European nation’s adjuvant use in influenza vaccines.

VII. THE QUEST FOR A ‘‘UNIVERSAL’’ INFLUENZA VACCINE: IS IT POSSIBLE? The quest to develop a ‘‘universal influenza vaccine’’ that protects against all influenza subtypes continues. Many experts predict that the key to a universal vaccine may lie in targeting conserved antigenic epitopes, not only in well-accepted antigenic candidates (HA), but those not traditionally considered in the protective response (M2 and NP). The influenza M2 protein is embedded in the virus membrane, mediates virus fusion, and

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contains a nine residue epitope in the N-terminal ectodomain (M2e) that is highly conserved among all influenza subtypes. M2 and M2e-based vaccines have demonstrated potent humoral and cell-mediated heterosubtypic immune responses against conserved epitopes in numerous animal studies (Du et al., 2010; Ellebedy and Webby, 2009; Rimmelzwaan and McElhaney, 2008). Recent Phase I clinical trials demonstrated that the M2-based vaccines are safe and immunogenic in humans (Schotsaert et al., 2009). An alternative approach is targeting the conserved epitopes that exist within the HA stalk domain. Several motifs, including the HA fusion peptide (HA2), are targeted by broadly reactive neutralizing antibodies that can be administered therapeutically (Ekiert et al., 2009; Prabhu et al., 2009; Sui et al., 2009). Recent advances in isolating human monoclonal antibodies may provide increased understanding into the molecular basis for recognition and escape that underlies the constant antigenic drift in influenza surface proteins. They may also provide evidence for lifelong persistence of immunity to some influenza viruses (Crowe, 2009). Finally, NP may be a viable vaccine target. NP is produced in abundance during infection, and NP epitopes are potent CTL targets, especially during secondary infection (or in response to vaccination). Because it is an internal protein and is critical in regulating virus replication through interaction with the RNA genome, it remains highly conserved among influenza isolates. One drawback to vaccination with NP is the necessity for processing via the internal pathway, ultimately leading to presentation on antigen complexes that stimulate virus-specific CTLs. To overcome this hurdle, DNA vaccines and viral vectors have been employed. Both DNA immunization and recombinant pox viruses expressing NP generate a memory CTL response that is able to protect against multiple influenza subtypes (Altstein et al., 2006; Ulmer et al., 1993). Ultimately, a truly universal influenza vaccine may require inclusion of multiple conserved epitopes to be successful. Recent studies combining HA, M2, and NP in either DNA or viral-vectored vaccines have shown promising immunogenicity and cross-clade protection against the HPAI H5N1 viruses in animal models (Poon et al., 2009; Rao et al., 2010).

VIII. CONCLUSIONS In conclusion, outbreaks of influenza A viruses will remain a public health threat. Given an aging population and ever-changing virus, we must continue to improve our vaccination strategies. Many new and exciting approaches hint that the vaccine production process and even the influenza vaccines themselves will advance over time. The successful production and distribution of a new vaccine in the face of a pandemic is encouraging and suggests that we are on the right track. However, only time will tell how society will respond to the next pandemic.

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for the 2009-10 influenza season: a multicentre, randomised controlled trial. Lancet 375:49–55. Wan, H., et al. (2008). Replication and transmission of H9N2 influenza viruses in ferrets: Evaluation of pandemic potential. PLoS ONE 3:e2923. Wang, C., et al. (1993). Ion channel activity of influenza A virus M2 protein: Characterization of the amantadine block. J. Virol. 67:5585–5594. Wang, X., et al. (2000). Influenza A virus NS1 protein prevents activation of NF-kappaB and induction of alpha/beta interferon. J. Virol. 74:11566–11573. WHO (2006). A review of production technologies for influenza virus vaccines, and their suitability for deployment in developing countries for influenza pandemic preparedness. http://www.who.int/vaccine_research/diseases/influenza/Flu_vacc_manuf_tech_report. pdf. WHO (2007). Use of Cell Lines for the Production of Influenza Virus Vaccines: An Appraisal of Technical, Manufacturing, and Regulatory Considerations. http://www.who.int/vaccine_research/diseases/influenza/WHO_Flu_Cell_Substrate_Version3.pdf. WHO (2009a). Pandemic (H1N1) 2009. http://www.who.int/csr/disease/swineflu/en/ index.html. WHO (2009b). Strategic Advisory Group of Experts on Immunization - report of the extraordinary meeting on the influenza A (H1N1) 2009 pandemic. Wkly. Epidemiol. Rec. 84:301–308. WHO (2009c). Pandemic influenza A (H1N1) 2009 virus vaccine—conclusions and recommendations from the October 2009 meeting of the immunization strategic advisory group of experts. http://www.who.int/csr/disease/swineflu/meetings/sage_oct_2009/en/ index.html. WHO (2010a). http://www.who.int/csr/disease/avian_influenza/country/cases_table_ 2010_04_21/en/index.html. WHO (2010b). Antigenic and genetic characteristics of influenza A(H5N1) and influenza A (H9N2) viruses and candidate vaccine viruses developed for potential use in human vaccines. http://www.who.int/csr/disease/avian_influenza/guidelines/h5n1virus/en/. Wong, J. P., et al. (2009). Activation of toll-like receptor signaling pathway for protection against influenza virus infection. Vaccine 27:3481–3483. Wright, P. F. (2008). Vaccine preparedness-are we ready for the next influenza pandemic? N. Engl. J. Med. 358:2540–2543. Wu, F., et al. (2009). The co-administration of CpG-ODN influenced protective activity of influenza M2e vaccine. Vaccine 27:4320–4324. Xing, Z., et al. (2010). Host immune and apoptotic responses to avian influenza virus H9N2 in human tracheobronchial epithelial cells. Am. J. Respir. Cell Mol. Biol. doi: 10.1165/ rcmb.2009-0120OC. Yen, H. L., and Webster, R. G. (2009). Pandemic influenza as a current threat. Curr. Top. Microbiol. Immunol. 333:3–24. Yewdell, J., and Garcia-Sastre, A. (2002). Influenza virus still surprises. Curr. Opin. Microbiol. 5:414–418.

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4 Innate Host Barriers to Viral Trafficking and Population Diversity: Lessons Learned from Poliovirus Julie K. Pfeiffer

Contents

Abstract

I. Introduction A. The benefit of poliovirus research in an era of eradication II. Poliovirus Route and Barriers A. Phase I: Gastrointestinal tract B. Phase II: Blood and peripheral tissues C. Phase III: CNS III. Effects of the Host Barriers on Viral Population Dynamics A. RNA virus mutations B. Fitness loss from excessive mutagenesis C. Fitness loss from limited viral population diversity IV. Summary Acknowledgments References

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Poliovirus is an error-prone enteric virus spread by the fecal–oral route and rarely invades the central nervous system (CNS). However, in the rare instances when poliovirus invades the CNS, the resulting damage to motor neurons is striking and often permanent.

Department of Microbiology, University of Texas Southwestern Medical Center Dallas, Texas, USA Advances in Virus Research, Volume 77 ISSN 0065-3527, DOI: 10.1016/S0065-3527(10)77004-4

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In the prevaccine era, it is likely that most individuals within an epidemic community were infected; however, only 0.5% of infected individuals developed paralytic poliomyelitis. Paralytic poliomyelitis terrified the public and initiated a huge research effort, which was rewarded with two outstanding vaccines. During research to develop the vaccines, many questions were asked: Why did certain people develop paralysis? How does the virus move from the gut to the CNS? What limits viral trafficking to the CNS in the vast majority of infected individuals? Despite over 100 years of poliovirus research, many of these questions remain unanswered. The goal of this chapter is to review our knowledge of how poliovirus moves within and between hosts, how host barriers limit viral movement, how viral population dynamics impact viral fitness and virulence, and to offer hypotheses to explain the rare incidence of paralytic poliovirus disease.

I. INTRODUCTION Poliovirus is a nonenveloped enteric RNA virus in the Picornaviridae family that has the ability to invade the central nervous system (CNS), despite the fact that entering the CNS has no apparent benefit for viral transmission. In the periphery, poliovirus can replicate in many cell types. However, in the CNS, poliovirus replication and subsequent damage is limited to motor neurons. Paralysis from motor neuron damage is often permanent. For this reason, poliovirus has an infamous reputation and shaped the public view of infectious diseases in the twentieth century. In reality, the chances of a person developing serious complications from poliovirus infection are exceedingly small—much smaller than the chances of being seriously injured in an accident. In the prevaccine era, it is likely that most individuals within an epidemic community were infected; however, only 4–8% of those infected exhibited any symptoms of disease, with most of these developing an abortive mild febrile illness and nothing more (Pallansch and Roos, 2001). A small subset of those with symptoms developed aseptic meningitis, which was generally self-limiting. Only 0.5% of infected individuals developed paralytic poliomyelitis. Nonetheless, poliovirus terrified the public and launched a massive research effort that rivals that of current human immunodeficiency virus (HIV) research. These efforts were rewarded with two outstanding vaccines— the Salk inactivated vaccine and the Sabin live-attenuated vaccine. Due to the overwhelming success of the Salk and Sabin vaccines, poliovirus is no longer a public health threat in developed countries. In 1988, the World Health Organization began a campaign to eradicate poliovirus from the planet by the year 2000, using the live-attenuated trivalent Sabin poliovirus

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vaccine. Although the eradication attempt has been largely successful, wildand vaccine-derived poliovirus cases are still being reported in developing countries (Arita et al., 2006; Roberts, 2006a,b). There are several reasons for this potential failure, including the high proportion of asymptomatic carriers, lack of complete vaccine coverage in politically unstable regions, low immunity among some populations despite multiple vaccinations, and reversion of the attenuated vaccine strains. Additional research and resources will be required to eradicate poliovirus.

A. The benefit of poliovirus research in an era of eradication Some might question the utility of poliovirus research in an era where the virus has been eradicated from developed countries and global eradication efforts are ongoing. Why study poliovirus in 2010 and beyond? 1. Aid the eradication campaign. Wild poliovirus and poliomyelitis remain endemic to several countries and are spreading. Until recently, wild poliovirus was circulating in just four nations, Afghanistan, Nigeria, Pakistan, and India; however, new cases have emerged in several other countries. Additionally, the viruses within the Sabin live-attenuated vaccine frequently revert attenuating mutations, occasionally resulting in cases of vaccine-associated paralytic poliomyelitis (Kew et al., 1981; Nkowane et al., 1987). These revertant viruses can circulate and cause additional disease. Understanding poliovirus evolution and limiting reversion of attenuating mutations will aid eradication efforts. 2. Serve as a model for vaccine development. A long-standing question is why multiple successful vaccines were readily developed for poliovirus when other viruses resist vaccine development. Perhaps by understanding how and why the poliovirus vaccines work, other vaccine endeavors will be more successful. 3. Serve as a model for other enteric viruses. Many viruses, including several medically important enteric viruses, are difficult to study because of limited replication in cell culture and animal models. The robust growth of poliovirus, paired with tools developed over a century of research, makes poliovirus an attractive model system for understanding the basic biology of other viruses. 4. Prepare for viruses that will occupy the niche vacated by poliovirus in the posteradication era. Humans have coexisted with poliovirus for millennia, in a largely stable, minimally pathogenic relationship. It is possible that poliovirus eradication will open a niche to be occupied by another pathogen. Likely candidates include other enteroviruses in the Picornaviridae family. Knowledge gained from poliovirus research will inform efforts to guard against future epidemics.

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5. Understand other enteroviruses. Poliovirus has lifestyle similar to other occasionally pathogenic enteroviruses. Enteroviruses include poliovirus, echovirus, coxsackievirus, and human enterovirus and cause 1 billion infections per year worldwide (Oberste et al., 2000). Apparent disease is relatively rare, but enteroviruses account for 5–10 million symptomatic infections per year in the United States (Strikas et al., 1986) and are the leading cause of the most common CNS infection, aseptic meningitis. Of particular concern are enterovirus 71 (EV71) and coxsackievirus B3 (CVB3). EV71 is the most significant neurotropic enterovirus in some areas of the world and is likely to be the most important neurotropic/ paralytic enterovirus upon eradication of poliovirus (Chumakov et al., 1979; McMinn, 2002; Pallansch and Roos, 2001; Shindarov et al., 1979). CVB3 can cause paralysis and myocarditis and may lead to the development of type I diabetes (Grist et al., 1978; Pallansch and Roos, 2001). Over 3% of CVB3 infections result in cardiac signs or symptoms (Grist et al., 1978). However, most enterovirus infections are asymptomatic; therefore, understanding how poliovirus moves within a host to initiate disease will shed light on the trafficking and disease course of other enteroviruses.

II. POLIOVIRUS ROUTE AND BARRIERS A. Phase I: Gastrointestinal tract 1. Poliovirus in the gastrointestinal tract In order to propagate itself, poliovirus must be shed from a primary host, survive in the environment, and initiate a productive infection in the gastrointestinal (GI) tract of a secondary host. Most infected individuals shed poliovirus in their feces for 2–8 weeks after infection (Nathanson, 2008). Intrafamily transmission is rapid and relatively complete (Pallansch and Roos, 2001). Many poliovirus infections are transmitted by direct person-to-person contact, without the need for long-term viral survival in the environment. However, poliovirus infections are also initiated by virus in the environment, via fomites, etc. Although the precise amount of virus required for natural infection is not known, experiments with the Sabin vaccine viruses suggested that 100 viable virions (tissue culture infectious dose units) can productively infect infants (Minor et al., 1981). After ingestion, poliovirus must bind to and enter cells via its cellular receptor, CD155 (also called PVR, for poliovirus receptor), which is a major susceptibility factor (Mendelsohn et al., 1989). While the normal function of CD155 is not completely understood, it is ubiquitously expressed in humans and is involved in the formation of adherens

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junctions and natural killer cell recognition (Racaniello, 2006). Importantly, not all tissues that express CD155 are targets for poliovirus replication, and some tissues with low CD155 expression, such as the intestine, are known sites of poliovirus replication (Bernhardt et al., 1994; Ren and Racaniello, 1992a; Ren et al., 1990; Solecki et al., 1997). The importance of human CD155 as a susceptibility factor is highlighted by the finding that mice are not susceptible to poliovirus infection unless they express a human CD155 transgene (Koike et al., 1991; Ren et al., 1990). CD155transgenic mice are susceptible to poliovirus infection and paralytic disease from injected virus; however, they are not susceptible by the natural oral route of infection. The initial explanation for the oral insusceptibility of CD155-transgenic mice was that CD155 expression was not detected in the GI tract (Koike et al., 1991; Ren et al., 1990). Several efforts to create orally susceptible mice by using alternate promoters to drive CD155 expression were unsuccessful (Crotty et al., 2002; Ida-Hosonuma et al., 2002; Yanagiya et al., 2003; Zhang and Racaniello, 1997), despite detectable expression of CD155 in intestinal tissue (Crotty et al., 2002; Yanagiya et al., 2003) and binding of virus to intestine fragments ex vivo (Zhang and Racaniello, 1997). These results indicate that CD155 expression is one factor of many required for murine infection by the oral route. After poliovirus enters the mouth, it may replicate in the oropharynx before transit to the lower GI tract (Bodian and Horstmann, 1965; Melnick, 1996; Sabin, 1956). The cell types infected in the oropharynx are not known but may include lymphatic tissues. While poliovirus undergoes robust replication in the oropharynx of chimpanzees (Sabin, 1956), replication in the human throat appears to be limited, and virus is rapidly cleared, despite long-term replication in the intestine (Bodian, 1952; Sabin and Ward, 1941b). These data imply that viral replication in the oropharynx may be limited and transient in humans. Productive infection requires viral survival in the stomach and access to and replication within susceptible cells in the intestine; however, the identity of the infected cell type remains unclear. The intestinal mucosa is composed of several different cell types. Epithelial cells, including enterocytes, Paneth cells, and goblet cells, create a barrier that limits microbial translocation while allowing absorption of nutrients from the lumen. Subepithelial immune cells such as intraepithelial lymphocytes and lamina propria lymphocytes, including IgA-producing B cells, play important roles in controlling commensal and pathogenic microorganisms. The intestinal mucosa contains regions of specialized lymphatic tissues called Peyer’s patches. Within Peyer’s patches, specialized M cells within the epithelium sample gut antigens and present them to underlying immune cells, which can circulate via the lymphatic system. Virus may initially enter and replicate in intestinal lymphatic tissues or adsorptive epithelial cells (enterocytes; Fig. 1). While there is evidence for viral infection of each

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Lumen Bacteria

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FIGURE 1 Architecture of the gastrointestinal epithelium. Virus may access the epithelium through M cells and the lymphatic system, or epithelial cells. IEL, intraepithelial lymphocyte; LPL, lamina propria lymphocyte; DC, dendritic cell; PP, Peyer’s patch.

cell type, more evidence appears to be in favor of initial infection via cells of the lymphatic system. First, poliovirus was visualized and isolated from human and chimpanzee lymphatic tissue (Bodian, 1955a; Bodian and Horstmann, 1965; Sabin and Ward, 1941a,b) as well as mouse lymphatic tissue following intraperitoneal injection (Buisman et al., 2003). Second, poliovirus injected directly into the ileum of rhesus macaques was visualized in M-like cells, the lamina propria, and within lamina propria macrophages (Takahashi et al., 2008). Third, poliovirus was visualized in M cells after incubation of virus with human tissue containing Peyer’s patches (Sicinski et al., 1990). Fourth, poliovirus was exocytosed through M-like cells in culture systems containing monolayer cultures with M cells from human Peyer’s patches (Ouzilou et al., 2002). Fifth, poliovirus can replicate in isolated lymphoid cells such as dendritic cells and macrophages, particularly after cellular activation (Eberle et al., 1995; Freistadt and Eberle, 1996; Freistadt et al., 1993; Wahid et al., 2005; Willems et al., 1969). Despite this evidence, it is unknown whether poliovirus actively replicates in lymphoid tissues in vivo, or whether it simply drains there. Evidence for viral replication in the superficial epithelium includes a poliovirus strain (Leon) that did not appear to replicate in lymphoid tissues but nonetheless replicated in the gut (Sabin, 1955, 1956) and

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visualization of poliovirus in microvilli epithelial cells, but not M cells, within the intestine of CD155-transgenic mice (Ohka et al., 2007). The cellular origin for viral shedding and transmission is also unclear. While virus may initially enter and replicate in intestinal lymphatic tissues, gut epithelial cells are candidates for viral shedding into the lumen (Nathanson, 2008). CD155 is localized primarily on the basolateral surface of polarized human CaCo-2 cells; therefore, poliovirus may access epithelial cells from the back, replicate, and be shed apically into the lumen (Boot et al., 2003; Ohka and Nomoto, 2001). This model is supported by data from the Cutter incident, where children were administered incompletely inactivated poliovirus by intramuscular injection. Perhaps predictably, several of these children developed paralytic poliomyelitis; however, contacts of these children were also infected with poliovirus, with 113 developing paralytic disease (Nathanson and Langmuir, 1963a,b; Offit, 2005). Since most of these secondary infections were through the fecal–oral route, virus was able to travel from the injection site to the intestinal tract of the vaccine recipient, with productive viral replication and shedding. Therefore, it is likely that virus accessed the gut lumen by entering the basolateral surface of the epithelium in the vaccine recipients. Experiments with human volunteers suggest that viral replication in the lower gut is required for productive shedding in feces (Sabin, 1956). Overall, it is possible that poliovirus entry and replication in the gut is similar to that of reovirus. Reovirus enters M cells overlying Peyer’s patches, enters intestinal epithelial cells from the basolateral surface, replicates, and is shed into the lumen (Bass et al., 1988; Excoffon et al., 2008; Rubin et al., 1985; Wolf et al., 1981, 1983). More work is required to identify the specific cell types infected by poliovirus in the gut.

2. Host barriers in the GI tract Poliovirus must navigate several hurdles to initiate productive infection in the gut. First, virus must survive the interhost period and resist inactivation from desiccation, alternate temperatures, and pH, etc. It is thought that poliovirus cannot survive in the environment for more than a couple of weeks, but under optimal conditions of neutral pH, sufficient moisture, and low temperature, the virus may survive for months (Pallansch and Roos, 2001; Tyler and Nathanson, 2001). Many instances of poliovirus transmission are through direct person-to-person contact rather than exposure through environmental sources such as sewage. These personto-person transfers are likely to be much more efficient than infection by exposure to environmental sources of virus. Once poliovirus enters the body through the mouth, several nonspecific host barriers are encountered. Saliva, mucus, and cilia may aid viral clearance prior to replication. Passage through the stomach may also pose a challenge to the virus (Salo and Cliver, 1976). The stomach pH is 2.0 or

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lower, and virus must survive stomach acid prior to reaching the small intestine, where pH ranges from 6.5 to 7.5 (Ewe et al., 1999). A recent study examined the sensitivity of poliovirus to stomach acid in a murine model and in vitro. The results suggest that poliovirus is inactivated by stomach acid after prolonged incubation. When virus was delivered to fasted mice by a gastric tube, little of the inoculum was found in small intestine, although recovery was aided if a pH-neutralizing agent was coadministered with the virus (Ohka et al., 2007). In a different study, CD155transgenic mice were orally inoculated with a pool of ten marked viruses to identify host barriers and viral bottleneck effects (Fig. 2). While all 10 viruses were often found in feces, only one or two marked viruses were found in the CNS (Kuss et al., 2008). These results suggest that the barrier encountered during transit to the CNS was much more severe than the barrier encountered during passage through the GI tract. Although the barrier imposed by the GI tract may be less severe than the barrier encountered during trafficking to the CNS, recent experiments from our laboratory suggest that only
1% of the total inoculum is shed in feces of orally inoculated mice (unpublished data; Kuss et al., 2008). Stomach acid is not the only barrier encountered in the upper GI tract; the virus must resist inactivation by bile salts and digestive enzymes. In general, enveloped viruses are susceptible to inactivation by bile salts, but nonenveloped viruses such as poliovirus are quite resistant. Proteases are secreted by gastric and pancreatic cells, and enteric viruses are either resistant to cleavage or use proteases to their advantage. For poliovirus, ex vivo exposure to proteases present in gastric contents did not reduce viral titer (Ohka et al., 2007). Viruses in the Reoviridae family are cleaved by intestinal proteases, which increase viral infectivity in vitro and in vivo (Amerongen et al., 1994; Bass et al., 1990; Clark et al., 1981; Estes et al., 1981), suggesting that proteases may actually aid some enteric viruses. Throughout the GI tract, a mucus layer lines the epithelium, and viruses must penetrate this layer to gain access to susceptible cell types below. Mucus overlying gut epithelia exists in two layers. The outer, loosely attached layer is 100 mm thick and the inner, firmly attached layer is 50 mm thick (Atuma et al., 2001). The inner layer is largely devoid of bacteria ( Johansson et al., 2008). Together, these layers form a 150 mm viscous barrier, which is 5000 times the 30 nm diameter of a poliovirus particle. By analogy, this layer would amount to a human swimming through 150 gel-filled American football fields. Several mucin genes have been identified, and mucin proteins are heavily glycosylated, forming a net-like structure (Dekker et al., 2002; Johansson et al., 2008). Mucins can inhibit enteric viruses such as rotavirus (Chen et al., 1993) and protect the host from bacterial pathogens such as Campylobacter (McAuley et al., 2007) and Citrobacter species (Bergstrom et al., 2010). The precise effect of the mucus layer on viral infection is unknown.

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FIGURE 2 Identification of poliovirus barriers using a viral population diversity assay. (A) Silent mutations in the capsid-coding region (indicated in bold underline) were used to genetically mark ten polioviruses such that host barriers and viral population bottlenecks could be identified in an orally susceptible poliovirus mouse model (CD155þ/þ IFNAR/). (B) Following infection and isolation of total RNA, the tag region was amplified by RT-PCR, blotted on a membrane, and probed with 32P-labeled oligonucleotides matching each tag sequence. (C) Each probe hybridized specifically with its cognate virus. (D) Samples were harvested from mice orally inoculated with the ten-virus pool upon disease onset, and data from the hybridization assay revealed host barrier-mediated viral population restriction during trafficking within the host. Data from two representative mice are shown. AA, amino acid; WT PV, wild-type poliovirus sequence; Sm. Int., small intestine. Data are from Kuss et al. (2008).

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In addition to the barrier function created by mucus, the mucus layer contains secretory IgA, and other secreted factors that may have antiviral activity. Over 75% of immunoglobulin in the body is IgA, and most is secreted across mucus membranes. IgA is important for homeostasis of the gut environment and is impacted by the gut microflora (Macpherson and Slack, 2007; Macpherson et al., 2008; Tomasi et al., 1965). For poliovirus, the appearance of poliovirus-specific IgA in the intestine coincides with decline in poliovirus shedding and eventual clearance (Valtanen et al., 2000). Therefore, IgA plays a role in control of poliovirus infections in the gut. As a member of the immunoglobulin superfamily, the PVR CD155 has the potential to limit viral infection through a secreted form of the receptor. There are four isoforms of human CD155; two isoforms are membrane bound and two isoforms are secreted (Baury et al., 2003). The secreted forms of CD155 are detectable in cultured cell medium, human serum, and human cerebrospinal fluid and have the capacity to limit viral entry by blocking binding to membrane-bound CD155 (Baury et al., 2003). In fact, many viral receptors are members of the immunoglobulin superfamily, and since most viruses bind these proteins at the most membrane-distal site, it has been suggested that soluble forms of viral receptors may be precursors to modern antibodies and that soluble receptors neutralize viruses by competing with receptors on the cell surface (Dermody et al., 2009). While this idea of soluble receptor-mediated poliovirus neutralization has not been examined in vivo and may be operative during human infections, current transgenic mouse models express only the membrane-bound form of CD155; therefore, soluble CD155 is unlikely to contribute to the major host barriers observed in this mouse model (see below; Koike et al., 1990, 1991; Kuss et al., 2008; Lancaster and Pfeiffer, 2010; Ren et al., 1990). A variety of relatively nonspecific host defense mechanisms may contribute to enteric virus barriers in the gut lumen. Defensins are hostderived antimicrobial proteins produced by specialized gut epithelial cells (Ayabe et al., 2000; Salzman et al., 2007). While defensins are generally thought of as antibacterial proteins due to their membrane disruption potential, there is evidence that defensins may target both enveloped and nonenveloped viruses (Buck et al., 2006; Daher et al., 1986; Gropp et al., 1999; Hazrati et al., 2006; Wang et al., 2004b). For nonenveloped viruses such as adenovirus, the antiviral activity of defensins is independent of membrane disruption and may involve inhibition during intracellular steps in the viral replication cycle (Smith and Nemerow, 2008). Lactoferrin is another relatively nonspecific host protein with antibacterial and antiviral properties. Lactoferrin is present in milk, mucosal secretions, and secretory granules of neutrophils and inhibits bacteria through sequestration of iron. Lactoferrin has antiviral activity for several viruses including rotavirus, enteroviruses, and adenovirus and is thought to act by inhibiting viral internalization (Seganti et al., 2004; Weng et al., 2005).

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Once a virus reaches the cell surface, it must enter and initiate steps to ensure propagation. It is uncertain which cell types are infected by poliovirus in the gut. If the initial entry event occurs in epithelial cells, it is likely to require expression of the PVR, CD155. Viral receptor expression in the intestinal epithelium, M cells, and germinal centers correlates with oral susceptibility (Iwasaki et al., 2002). For poliovirus, humans are the most orally susceptible, followed by chimpanzees, Old World monkeys, New World monkeys, and CD155-transgenic mice (Iwasaki et al., 2002; Sabin, 1956). While CD155 expression is barely detectable or undetectable in several strains of non-orally susceptible CD155-transgenic mice (Koike et al., 1991; Ren et al., 1990), viral replication can still occur in the gut and replicated virus is shed in feces (our unpublished data; Boot et al., 2003; Kuss et al., 2008), suggesting that other host factors contribute to the lack of disease in these mice following oral inoculation (see discussion below on the role of type I interferon, etc.). If the initial entry event occurs via trancytosis through M cells, the process may be CD155 dependent or CD155 independent. M cells sample gut antigens nonspecifically, so it is possible that viral entry and trancytosis through M cells may not require the viral receptor. However, poliovirus replication requires CD155, since viral uncoating is triggered by receptor engagement. Studies suggest that binding and entry in M cells may be very inefficient, since less than 1% of the input virus was trancytosed through M cells in a cell culture model (Ouzilou et al., 2002), and the proportion of poliovirus bound to M cells in ex vivo human Peyer’s patch tissue was minimal, even compared with reovirus in a similar model (Sicinski et al., 1990; Wolf et al., 1981). Overall, the process of viral entry into gut tissues is likely to be inefficient, constituting a major barrier for viral replication. The host’s innate immune response through induction of interferons and interferon-induced genes is a major impediment to viruses. Viral replication generates a variety of pathogen-associated molecular pattern molecules, such as double-stranded RNA, that are detected by the host and lead to induction of a variety of antiviral effectors. Key components of this pathway are type I interferons, interferon a and b, which are produced by many cells types in response to viral infection. IFNa/b are released from infected cells and bind the interferon a/b (IFNAR) receptor on neighboring cells, which are protected by an induced antiviral state. The importance of IFNAR in limiting many viruses has been demonstrated in experiments with mice lacking the IFNAR gene (Fiette et al., 1995; Garcia-Sastre et al., 1998; Mrkic et al., 1998; Muller et al., 1994; Ryman et al., 2000; Samuel and Diamond, 2005; Shresta et al., 2004; Wessely et al., 2001). IFNAR knockout mice demonstrate increased susceptibility to a wide variety of viruses, including poliovirus. In fact, CD155-transgenic mice are not orally susceptible unless they lack IFNAR expression, indicating the importance of the type I interferon pathway in protecting the

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host (Ida-Hosonuma et al., 2005; Ohka et al., 2007). Interestingly, poliovirus can replicate in the gut of CD155 mice that have functional IFNAR, suggesting that the inhibitory effect of interferon may act at a postgut step (see below; Boot et al., 2003; our unpublished data). The role of IFNa/b in limiting viral replication in the periphery will be discussed in more detail in the next section. Recently, type III interferon (also known as IFNl or IL-28/29) has emerged as an important contributor to host innate immune responses, particularly at mucosal surfaces (Kotenko et al., 2003; Sheppard et al., 2003). The response of IFNl to infection with several viruses, including Theiler’s murine encephalitis virus and mouse hepatitis virus, is particularly high in stomach, intestine, and lung and appears to protect epithelial surfaces (Mordstein et al., 2010; Sommereyns et al., 2008). The gut architecture and natural homeostasis may be another obstacle to enteric viruses. Epithelial cells are constantly sloughing and are being shed from villi. Therefore, enteric viruses using epithelial cells for replication and shedding must constantly infect new cells to maintain prolonged shedding (Boshuizen et al., 2003). Cells of the GI tract are joined by tight junctions that prevent passage of pathogens and commensals. Viruses requiring access to lymphatics or blood must penetrate this impressive barrier. Using a pool of ten marked polioviruses, it was observed that most viruses in the gut lumen are unable to access gut tissues or pass the gut barrier and enter the bloodstream and/or CNS (Kuss et al., 2008). However, if the gut epithelium was disrupted with dextran sulfate sodium treatment prior to oral poliovirus infection, more of the marked viruses were observed in gut tissues and blood. Interestingly, gut damage was unable to allow more viruses in the population to reach the CNS, suggesting that the gut epithelium is one barrier of several which must be overcome for poliovirus transit to the CNS (Kuss et al., 2008).

B. Phase II: Blood and peripheral tissues 1. Poliovirus in blood and peripheral tissues Following primary viral replication in the mucosa, it is thought that poliovirus drains into cervical and mesenteric lymph nodes, and then into blood. Virus is only detectable in the blood if the rate of viral input due to replication exceeds the rate of clearance by the reticuloendothelial system and other mechanisms. Transient viremia is common during poliovirus infections, and this primary viremia may occur in most people and orally infected chimpanzees (Bodian, 1952; Horstmann et al., 1954a,b; Melnick et al., 1961). A subset of infected individuals, roughly 4–8%, develop a secondary, major viremia due to continued viral replication in extraintestinal sites (Bodian and Horstmann, 1965; Bodian and Paffenbarger, 1953; Horstmann et al., 1954a,b; Melnick, 1996; Sabin,

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1956). The secondary, major viremia is often accompanied by symptoms of minor illness and is a prerequisite for the development of CNS disease. At this stage, virus is thought to replicate in non-neural sites including brown fat, reticuloendothelial tissue, and muscle (Bodian, 1955a; Ren and Racaniello, 1992a; Sabin, 1956; Wenner and Kamitsuka, 1957).

2. Host barriers in blood and peripheral tissues The three major barriers limiting poliovirus during the blood and peripheral tissue stage are clearance by the reticuloendothelial system, innate immune responses, and adaptive immune responses. Even prior to extraintestinal replication or development of an adaptive immune response, virus is cleared by phagocytic cells of the reticuloendothelial system. For other viruses, such as yellow fever virus and herpes simplex virus, inhibition of macrophage phagocytosis increases viremia (Irie et al., 1998; Tyler and Nathanson, 2001). Phagocytic clearance is impacted by the size of the particle; therefore, opsonization of viruses with antibody or complement increases efficiency. Innate immune responses are critical components of the host’s defense against viral infections, and for poliovirus, the innate immune response primarily limits viral replication during the blood–extraintestinal tissue phase. Poliovirus and other enteric picornaviruses are sensitive to type I interferon in vivo (Fiette et al., 1995; Ida-Hosonuma et al., 2005; Wessely et al., 2001; Yoshikawa et al., 2006). As described above, CD155 transgenic mice are not orally susceptible to poliovirus unless they have limited IFNa/b responses due to IFNAR gene deletion (Ohka et al., 2007). Additionally, poliovirus replication is enhanced in CD155-expressing-IFNAR knockout mice, with viral replication and high viral titers in tissues, such as liver, pancreas, and kidney, which are not normally sites of replication in immune-competent mice (Ida-Hosonuma et al., 2005; Yoshikawa et al., 2006). Poliovirus susceptibility in various tissues correlates with amount of interferon induction and levels of interferon-stimulated genes prior to infection (Ida-Hosonuma et al., 2005; Ohka et al., 2007; Yoshikawa et al., 2006). These results prompted Ida-Hosonuma et al. (2005) to propose that visceral organs of immune-competent mice, which are more likely to encounter microbes leaking from the intestinal tract, are primed for a fast response to infection, while CNS tissues are not. While a pool of ten marked polioviruses was severely restricted during travel to the CNS of immune-competent CD155-transgenic mice after intraperitoneal or intramuscular injection, this restriction was dramatically diminished in CD155-IFNAR knockout mice (Kuss et al., 2008; Lancaster and Pfeiffer, 2010). Taken together, these studies highlight the importance of innate immune responses, particularly type I interferon, in restricting poliovirus spread in the periphery.

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The best-known barrier limiting poliovirus viremia and the development of disease is neutralizing antibody. Decades of research have shown that neutralizing antibody protects humans and primates from paralytic disease by preventing viremia (Nathanson, 2008). Passive antibody delivered prior to infection is protective, and there is a strong correlation between poliovirus antibody titer and protection from paralysis (Bodian, 1953, 1955a; Sabin, 1956). However, serum antibodies do not prevent viral replication in the gut, since humans vaccinated with inactivated virus via injection still allow replication of the live-attenuated vaccine after oral delivery (Horstmann, 1955; Sabin, 1956). Children vaccinated with the live-attenuated vaccine become infected after challenge with a second dose, but they shed less virus over a shorter time course, suggesting that poliovirus-specific IgA may limit viral replication in the gut (Horstmann, 1955; Sabin, 1956). During primary infections, clearance of viremia is immediately followed by detectable serum antibody specific for poliovirus (Nathanson, 2008). The efficacy of the inactivated/injected poliovirus vaccine suggests that serum IgG is sufficient to prevent paralysis; however, the additional mucosal IgA induced by the live-attenuated/ oral vaccine may at least partially limit viral replication in the gut and subsequent shedding. The dramatic success of the Salk and Sabin poliovirus vaccines is proof that neutralizing antibodies constitute a major host barrier to viral replication, spread, and CNS invasion.

C. Phase III: CNS 1. Poliovirus access to the CNS Poliovirus rarely invades the CNS, either through the blood–brain barrier or trafficking in neurons. CNS infection and subsequent damage is an accidental diversion from the GI tract, with apparently little benefit to the virus. While poliovirus can undergo robust replication in the CNS and generate high viral titers, CNS virus is not likely to be transmitted to a new host by the normal fecal–oral route. The route of viral CNS entry has been debated for decades, with evidence for both blood and neural transit routes. Regardless of the CNS entry mechanism, it has been established that viremia must precede viral CNS invasion.

a. Poliovirus access to the CNS via the blood–brain barrier The blood– brain barrier is composed of microvasculature containing tight junctions with dense basement membranes, such that an active transport or carrier system is required for passage through these specialized capillaries. Some viruses, including West Nile virus and adenovirus, breach the blood– brain barrier to access the CNS (Dai et al., 2008; Gralinski et al., 2009; Wang et al., 2004a, 2008). This blood–brain barrier disruption can be observed by influx of dyes normally sequestered from the CNS.

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Poliovirus infection does not appear to induce blood–brain barrier disruption (Yang et al., 1997), and Evan’s blue dye does not enter the brain in murine infection models (our unpublished data). Yang et al. (1997) examined the ability of poliovirus to enter the CNS via the blood route using 35 S-labeled virions in intravenously injected mice. Poliovirus was observed in many tissues, and although the overall amount of virus in the cerebrum and cerebellum was the lowest, it was much greater than the theoretical amount predicted based on the vascular volume of the brain. Importantly, the amount of labeled virus in the brain at 5 h postinfection was similar in CD155-transgenic and -nontransgenic mice, suggesting that brain access was viral receptor-independent (Yang et al., 1997). However, in this study, the brains were not perfused prior to radioactive counting, and due to the high particle:plaque-forming unit ratio of poliovirus, it is uncertain whether viruses found in the brain are functional. Nonetheless, this study provided precedent that poliovirus entry into the CNS via the blood route can be modeled in mice. Poliovirus may use a variety of mechanisms to access the CNS through the blood–brain barrier. First, the CNS contains regions with reduced blood–brain barrier strength. In these sites, such as the area postrema in medulla oblongata, the capillary endothelium lacks tight junctions, which may allow more efficient transport of viruses (Bodian, 1952; Tyler and Nathanson, 2001). Second, it has been proposed that picornaviruses can directly infect endothelial cells (Blinzinger et al., 1969; Couderc et al., 1990; Friedman et al., 1981; Zurbriggen and Fujinami, 1988). Third, virus entry into the CNS may be aided by factors that increase vascular permeability (Sellers, 1969). Fourth, some viruses, such as measles and mumps, use transendothelial transport via diapedesis of immune cells. A ‘‘Trojan horse’’ mechanism for poliovirus transport in macrophages has been proposed, but not yet proven experimentally (Freistadt et al., 1993).

b. Poliovirus access to the CNS via neurons Like many other viruses, poliovirus can infect neurons and undergo axonal transport. Infection of neurons and transport mechanisms have been investigated for a variety of viruses including herpesviruses (Card et al., 1991; Smith et al., 2001), rabies (Iwasaki and Clark, 1975), reovirus (Morrison et al., 1991), West Nile virus (Samuel et al., 2007), adenovirus (Salinas et al., 2009), and picornaviruses (Martinat et al., 1999). Early evidence for poliovirus infection of neurons was generated in primate studies in the 1930s, 1940s, and 1950s (Bodian and Howe, 1940; Faber et al., 1953; Fairbrother and Hurst, 1930; Howe and Bodian, 1942; Verlinde et al., 1955). Poliovirus was found in nerve fibers after intramuscular, intravenous, or oral infection of monkeys. However, the role for viral transport in neurons was debated since nerve freezing or transection blocked viral transport in some studies, but not others (Bodian, 1954; Howe and Bodian, 1941; Nathanson and Bodian, 1961;

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Racaniello, 2006). These and other data sparked debate about the mechanism of poliovirus CNS entry, which is still not completely understood today. CD155-transgenic mice have been a useful tool for examining poliovirus neural transport mechanisms. Unlike in monkeys, in CD155-transgenic mice, there is consensus that poliovirus travels to the CNS via the neural route after intramuscular injection. In this mouse model, the injected limb is always the first limb paralyzed, and sciatic nerve transection protects mice from paralytic disease following intramuscular injection in the leg (Ohka et al., 1998; Ren and Racaniello, 1992b). It is thought that virus enters the sciatic nerve at the neuromuscular junction (Ohka et al., 2004), travels up the sciatic nerve to the spinal cord, and is then transported to the brain. Sciatic nerve transection experiments have revealed that poliovirus is transported by the fast retrograde axonal transport system, at a rate of over 12 cm/day (Ohka et al., 1998; Ohka et al., 2009). Transport in the sciatic does not require replication in muscle (Lancaster and Pfeiffer, 2010; Ohka et al., 1998), and intact 160S viral particles are observed in the sciatic nerve (Arita et al., 1999; Ohka et al., 1998, 2009). Perhaps not surprisingly, poliovirus does not undergo replication in long axons of the sciatic nerve (Lancaster and Pfeiffer, 2010). The cytoplasmic tail of the viral receptor CD155 interacts with Tctex-1 (Tctel-1), a light chain of cytoplasmic dynein (Mueller et al., 2002; Ohka et al., 2004). Taken together, these data suggest a model where poliovirus enters neurons at the neuromuscular junction via receptor-mediated endocytosis, the cytoplasmic tail of the receptor binds the dynein motor complex, and the virus-containing endosome is transported on microtubules to the cell body via the fast retrograde axonal transport system (Fig. 3). Uncoating and replication occur in the neural cell body, which can be very distant from the viral point of entry at the neuromuscular junction. Poliovirus entry and uncoating in neurons is very different from that in HeLa cells, where viral uncoating takes place in close proximity to the plasma membrane (Brandenburg et al., 2007). Therefore, neurons represent a unique environment, very distinct from cells found in the gut and periphery.

2. Host barriers en route to the CNS Regardless of whether poliovirus enters the CNS via a blood or neural route, several barriers may limit viral access. Poliovirus variants found in the CNS of human vaccine-associated paralytic poliomyelitis patients corresponded to only a small subset of viruses found in the gut, suggesting a barrier limited CNS access (Georgescu et al., 1994, 1997). Similarly, when CD155-transgenic mice were orally inoculated with a pool of ten marked polioviruses, only a small subset was found in the CNS (Kuss et al., 2008; Lancaster and Pfeiffer, 2010). Therefore, regardless of the

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Retrograde axonal transport Cell body Axon

Microtubule

Dynein complex CD155

FIGURE 3 Model for poliovirus trafficking in neurons. Poliovirus engages CD155 at the nerve terminal and is endocytosed. The cytoplasmic tail of CD155 interacts with a component of the dynein complex, and the virus-containing endosome moves through retrograde axonal transport to the cell body. Once in the cell body, viral uncoating and replication occur.

mechanism of CNS entry, following oral infection, viruses found in the CNS represent a small subset of the peripheral population, suggesting the existence of one or multiple host barriers limiting CNS access. Several viruses disrupt the blood–brain barrier, suggesting that it can impede viral transit in the absence of viral intervention (Soilu-Hanninen et al., 1994; Toborek et al., 2005; Wang et al., 2004a). In fact, a non-neuroinvasive variant of West Nile virus can enter the brain only if the blood– brain barrier was artificially breached (Kobiler et al., 1989). Despite the fact that poliovirus enters the CNS of CD155-transgenic mice from the blood (Yang et al., 1997), there is no evidence to suggest that poliovirus disrupts the blood–brain barrier. Therefore, the blood–brain barrier may impede poliovirus access to the CNS. Viral trafficking to the CNS via the neural route is subject to several barriers. First, viral infection can induce innate immune responses in neurons. Several viruses, including West Nile virus, herpesviruses, and rabies virus, induce production of type I interferon and other innate immune effectors in neurons (Delhaye et al., 2006; Griffin, 2003; Prehaud et al., 2005; Samuel et al., 2006). Second, substrates for viral replication are limited in axons. Poliovirus replication requires ribosomes, membrane vesicles derived from intracellular membrane compartments, and other factors that are unlikely to be plentiful in long axons or even nerve terminals. Therefore, viral replication is likely to be limited to neural cell bodies. Two lines of evidence suggest that poliovirus replication does not occur in long axons using a sciatic nerve model in CD155-transgenic mice:

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the presence of intact 160S particles (Arita et al., 1999; Ohka et al., 1998, 2009) and lack of viral replication scored by dye-containing virions (Lancaster and Pfeiffer, 2010). Third, viral trafficking in long axons is inefficient. Using a pool of marked polioviruses in a CD155-transgenic mouse model, only a subset of the input population was observed in the CNS following intramuscular inoculation (Kuss et al., 2008; Lancaster and Pfeiffer, 2010; Pfeiffer and Kirkegaard, 2006). Sciatic nerve sectioning revealed that, of the input population injected into the leg muscle,
80% was in the lower section of the sciatic nerve,
50% was in the middle section of the sciatic nerve, and only
20% was in the upper section of the sciatic nerve (Lancaster and Pfeiffer, 2010). These results suggest that retrograde axonal transport of poliovirus is inefficient, which may contribute to the low disease incidence from poliovirus infections. Studies with cultured neurons may aid our understanding of poliovirus transport barriers in neurons. However, a recent study demonstrated that cultured neurons do not completely recapitulate all transport mechanisms observed in mouse models (Ohka et al., 2009). Clues to CNS transport mechanisms and potential barriers have come from the phenomenon of ‘‘provocation poliomyelitis,’’ wherein increased poliovirus CNS disease follows peripheral tissue damage (McCloskey, 1950; Strebel et al., 1995). This tissue damage can be in the form of physical exertion, trivial muscle injury, or injections near the time of poliovirus infection. For example, trauma increased the incidence of paralysis in the injured limb, in naturally infected humans as well as in primate and mouse models (Bodian, 1954, 1955a; Gromeier and Wimmer, 1998; Hill and Knowelden, 1950; Lancaster and Pfeiffer, 2010; Ohka et al., 1998; Ren and Racaniello, 1992b). Similarly, injections administered within 30 days of the oral attenuated poliovirus vaccine or during poliovirus epidemics increased the incidence of paralytic disease (Anderson and Skaar, 1951; Guyer et al., 1980; Hill and Knowelden, 1950; McCloskey, 1950; Strebel et al., 1995). The ‘‘provoking’’ effect could be acting through a variety of mechanisms including localized alteration of the blood–brain barrier at corresponding areas of the CNS (Bodian, 1954, 1955a, 1955b; Sutter et al., 1992), inflammation, upregulation of the viral receptor (Nieke and Schachner, 1985; Squitti et al., 1999), increased viral replication, or increased efficiency of retrograde axonal transport (Gromeier and Nomoto, 2002). Peripheral injury may increase permeability of the corresponding CNS region (Trueta, 1955), but this mechanism has never been experimentally validated for poliovirus CNS access. There is evidence to suggest that muscle damage increases the efficiency of poliovirus retrograde axonal transport. Needle sticks delivered to intramuscularly injected CD155-transgenic mice increased the proportion of ten marked viruses able to traffic to the upper sciatic nerve and spinal cord (Lancaster and Pfeiffer, 2010). This muscle damage also increased the retrograde

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axonal transport efficiency of a nonviral protein, suggesting a general mechanism that may be involved in sensing tissue damage by relaying information to and from the cell body (Lancaster and Pfeiffer, 2010). The fact that tonsillectomy increased the incidence of bulbar/brainstem poliomyelitis supports the idea that damage to human tissue may increase neural transport of poliovirus (Modlin, 1995; Mueller et al., 2005). Overall, provocation poliomyelitis highlights the inefficiency of poliovirus CNS access in the absence of damage. Figure 4 contains a summary of poliovirus trafficking routes and potential host barriers discussed in this chapter.

Mouth

Ig

Heat, Desiccation, etc.

Ig

Mucus

Mucus

Virus in feces

Acid, etc.

Oropharyngeal mucosa

Intestinal mucosa Lumen factors, gut wall

Ig

Ig Inefficient retrograde axonal transport

ENS

Blood Ig

Ig

IFN

Skeletal muscle

BBB

CNS

IFN

Inefficient retrograde axonal transport

FIGURE 4 Summary of poliovirus trafficking routes and potential host barriers. In most infections, virus follows the course indicated by the bold arrows, entering the mouth, replicating in the intestinal mucosa, with shedding in feces. Virus can also enter the blood. Rarely, virus accesses the CNS through blood or neural routes. Potential host barriers limiting viral movement at each stage are indicated by gray boxes. BBB, blood– brain barrier; IFN, interferon; Ig, immunoglobulin; CNS, central nervous system; ENS, enteric nervous system (i.e., vagus nerve).

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III. EFFECTS OF THE HOST BARRIERS ON VIRAL POPULATION DYNAMICS Host barriers limit viral population diversity, and viral population diversity is hypothesized to be important for viral fitness and virulence. Therefore, in addition to their established role in limiting viral movement and replication, host barriers may also limit viral fitness through population bottleneck effects. This section describes the effect of host barriers on viral population dynamics and fitness.

A. RNA virus mutations The RNA-dependent RNA polymerases (RdRps) of poliovirus and many other RNA viruses are error-prone, giving rise to the highest mutation rates known in nature (Domingo and Holland, 1997; Drake, 1993). Observed error rates vary for different viruses and assays, but most measurements place the error frequency of RNA viruses at approximately one error per 10,000 nucleotides copied (Domingo and Holland, 1997; Domingo et al., 1978, 1996; Drake et al., 1998; Mansky and Temin, 1995). This translates into approximately one error per genome per cycle of replication. The errors incorporated by viral RdRps can have several consequences for the virus: the vast majority of mutations are deleterious, some mutations are neutral, and occasionally, under the right selective conditions, a few mutations are beneficial. Under selective pressure, mutations conferring resistance to neutralizing antibodies (van Doorn et al., 1995), cytotoxic T cell epitopes (Timm et al., 2004; Weiner et al., 1995), and antiviral drugs (Mansky, 2002) can emerge and take over the viral population, despite often having lower fitness than the parent virus in the absence of the selective pressure. Additionally, mutations that aid trafficking within the host may be required for virulence. RNA viruses are often referred to as ‘‘quasispecies,’’ a theoretical concept developed by Eigen and Schuster to describe complex populations (Eigen, 1996). Under certain conditions, RNA viruses are the best real-world example of the theoretical quasispecies (Domingo and Holland, 1997), and any description of RNA virus pathogenesis must take into account their incredible sequence diversity. A long-standing hypothesis in population genetics is that RNA viruses have evolved optimal error frequencies and that too many or too few errors will reduce the fitness of the viral population. Fitness is a measure of the ability to produce infectious progeny in a given environment (Domingo and Holland, 1997). Because of their high error rates, RNA viruses exist near the ‘‘error threshold’’: the value of fidelity where genetic information is irreversibly lost due to mutation, resulting in reduced fitness (Eigen, 1971). Presumably, RNA viruses risk over-mutagenesis of

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part of the population to reap potential benefits of enhanced adaptability for another subset of the population (Domingo and Holland, 1997).

B. Fitness loss from excessive mutagenesis Mutagen passage experiments have provided evidence for fitness loss from over-mutagenesis. Because of their high error rates, RNA viruses are very susceptible to mutagens, which extinguish the population through ‘‘error catastrophe’’ (Crotty et al., 2000; Domingo and Holland, 1997). In fact, the entire antiviral effect of the nucleoside analog drug ribavirin, at least for poliovirus, is due to enhanced mutagenesis of the viral genome (Crotty et al., 2001). Other viruses such as vesicular stomatitis virus, HIV, foot and mouth disease virus (FMDV), and lymphocytic choriomeningitis virus also undergo lethal mutagenesis upon mutagen treatment (Crotty et al., 2000; Holland et al., 1990; Loeb et al., 1999; Sierra et al., 2000). Mutagen-driven extinction of RNA viruses is a promising antiviral strategy; however mutagen-resistant isolates can emerge (Arnold et al., 2005; Pfeiffer and Kirkegaard, 2003; Sierra et al., 2007). Poliovirus passaged in the presence of ribavirin develops resistance to ribavirin and other mutagens via a single amino acid change, G64S, in the polymerase that increases the fidelity of RNA replication (Arnold et al., 2005; Pfeiffer and Kirkegaard, 2003). The complete crystal structure of the poliovirus RdRp revealed mechanistic insights into the enhanced fidelity of this polymerase: the mutation may alter the conformation of the active site, allowing more time for correct nucleoside triphosphate discrimination (Arnold et al., 2005; Thompson and Peersen, 2004).

C. Fitness loss from limited viral population diversity Pathogenesis experiments with high-fidelity, ribavirin-resistant G64S poliovirus have provided evidence for fitness loss due to low diversity (Pfeiffer and Kirkegaard, 2005; Vignuzzi et al., 2006). Although no growth defects were observed for the high-fidelity virus in cell culture, this virus was attenuated in mice (Pfeiffer and Kirkegaard, 2005). The attenuation was at least partly due to the increased fidelity of this virus, since it was less able to revert an additional attenuating mutation to restore virulence under selective pressure during replication in mice (Pfeiffer and Kirkegaard, 2005). Vignuzzi et al. (2006) showed the virulence of the high-fidelity virus could be restored by prepassage in mutagen to restore the quasispecies before murine inoculation. Viruses with high-fidelity RdRps show promise as live-attenuated vaccines (Vignuzzi et al., 2008). Together, these results suggest that viral population diversity is important for pathogenesis in infected animals, where a variety of selective pressures act on the viral population.

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Fitness can also be lost by fixation of deleterious mutations via genetic bottlenecks, and much has been learned about fitness limitations imposed by experimentally applied bottlenecks during viral passage in cell culture. For example, when RNA virus quasispecies diversity is repeatedly limited by in vitro serial plaque-to-plaque transfer bottlenecks, the resulting populations are less fit than the parental populations (Chao, 1990; Clarke et al., 1993; Duarte et al., 1992; Escarmis et al., 1996; Yuste et al., 1999). The amount of fitness loss is dependent on the fitness of the initial population (Domingo et al., 1978; Duarte et al., 1994). The bottleneck-imposed fitness loss correlates with the loss of the consensus, most-fit ‘‘wild-type’’ master sequence. This phenomenon is called ‘‘Muller’s ratchet’’: when small populations have high mutation rates and most mutations are deleterious, ‘‘a kind of irreversible ratchet mechanism’’ reduces fitness (Muller, 1964). Fitness can sometimes, but not always, be restored by passage at high multiplicities of infection (Duarte et al., 1993, 1994; Novella et al., 1995). Additionally, recombination-mediated genome repair can rescue populations from Muller’s ratchet (Felsenstein, 1974; Nee, 1988). Muller’s ratchet may be important for disease severity when bottlenecks exist during spread to, or spread within, hosts. Virus populations experience major bottleneck events during intrahost and interhost spread, and these events can impact the fitness and virulence of the remaining viral population. For example, in serial transmission events in pigs, FMDV experienced intrahost and interhost bottlenecks and the virus population lost virulence by the 14th passage (Carrillo et al., 1998, 2007). GB virus B, a relative of hepatitis C virus (HCV), experienced bottlenecks within experimentally infected primates, which may have reduced viral fitness (Weatherford et al., 2009). HIV and HCV undergo major bottlenecks during human transmission, with estimates of one or a few variants transmitted to the new host (Haaland et al., 2009; Laskus et al., 2004; McNearney et al., 1992). HIV spread within a host may also be restricted, since limited populations were observed in splenic germinal centers (Gratton et al., 2000). Despite these major bottleneck events, HIV and HCV quickly restore population diversity and remain highly successful pathogens. Poliovirus populations also experience bottlenecks in mouse models and in humans. Using a pool of ten marked viruses, the poliovirus population was severely restricted in the brain following inoculation by a variety of routes (intramuscular, intravenous, intraperitoneal, and oral; Kuss et al., 2008; Lancaster and Pfeiffer, 2010; Pfeiffer and Kirkegaard, 2006). Importantly, virulence, as measured by length of time to disease onset, correlated with viral population diversity in the brain. Mice with a more diverse viral population in the brain developed symptoms of disease significantly faster than mice with a less diverse population in the brain (Kuss et al., 2008). Overall, these studies suggest that poliovirus

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populations experience dramatic bottleneck effects due to host barriers. A similar stochastic bottleneck effect is seen in humans with vaccinederived paralytic poliomyelitis. Analysis of matched samples from stool and cerebrospinal fluid revealed that the CNS virus is just one member of the diverse population found in the gut (Georgescu et al., 1997). Interestingly, the most neurovirulent virus present in the gut was not always the ‘‘winner’’ found in the CNS (Georgescu et al., 1994, 1997). When inoculated into mice, the human vaccine-derived strains resulted in stochastic CNS invasion, typical of the bottleneck. Because a diverse viral population is required for full virulence, barriers that restrict viral quasispecies may reduce pathogenesis by limiting viral fitness in addition to limiting CNS entry (Domingo and Holland, 1997; Farci et al., 2002; Kimata et al., 1999; Kuss et al., 2008; Pfeiffer and Kirkegaard, 2005; Vignuzzi et al., 2006).

IV. SUMMARY Despite 100 years of poliovirus research, many questions remain unanswered. The rare incidence of paralytic poliomyelitis may be at least partially explained by barriers that limit viral trafficking within an infected host. Physical, anatomical, physiological, and immunological host barriers limit poliovirus replication, trafficking, and the maintenance of viral population diversity, all of which are required for virulence. Despite these barriers, poliovirus transmission persists, suggesting that the virus has evolved mechanisms to cope with countermeasures of the host. Because host barriers are operative for many viral and bacterial pathogens, understanding host barriers will aid not only the poliovirus eradication initiative but also efforts targeting other pathogens.

ACKNOWLEDGMENTS I thank the members of my laboratory for stimulating discussions and their contributions to this field. I also thank Chris Etheredge for preparing Figs. 1 and 3, and Gavin Best, Sharon Kuss, and Karen Lancaster for comments on the chapter. Work in the author’s laboratory is supported by NIH grant AI74668 and the Pew Scholars Program.

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5 Involvement of the Plant Nucleolus in Virus and Viroid Infections: Parallels with Animal Pathosystems M. E. Taliansky,* J. W. S. Brown,*,† M. L. Rajama¨ki,‡ J. P. T. Valkonen,‡ and N. O. Kalinina§

Contents

I. Introduction II. Structure and Functions of the Nucleolus III. What we have Learned about Interactions of Viruses with the Nucleolus from Animal Virology A. Viruses that replicate in the nucleus B. Viruses replicating in cytoplasm IV. Nucleolar Functions of Plant Virus Proteins and Viroids A. Nucleolus in replication of viroids and viruses B. The nucleolus and plant virus movement C. Nucleolar targeting for interference with host antiviral defense D. Nucleolar localization of viral proteins for as yet unknown reasons E. Formation of viral ribonucleoprotein complexes (RNPs) in the nucleolus V. Conclusions and Perspectives Acknowledgments References

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* Scottish Crop Research Institute, Invergowrie, Dundee, United Kingdom { { }

Plant Sciences Division, University of Dundee, United Kingdom Department of Agricultural Sciences, University of Helsinki, Helsinki, Finland A.N. Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia

Advances in Virus Research, Volume 77 ISSN 0065-3527, DOI: 10.1016/S0065-3527(10)77005-6

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2010 Elsevier Inc. All rights reserved.

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Abstract

The nucleolus is a dynamic subnuclear body with roles in ribosome subunit biogenesis, mediation of cell-stress responses, and regulation of cell growth. An increasing number of reports reveal that similar to the proteins of animal viruses, many plant virus proteins localize in the nucleolus to divert host nucleolar proteins from their natural functions in order to exert novel role(s) in the virus infection cycle. This chapter will highlight studies showing how plant viruses recruit nucleolar functions to facilitate virus translation and replication, virus movement and assembly of virus-specific ribonucleoprotein (RNP) particles, and to counteract plant host defense responses. Plant viruses also provide a valuable tool to gain new insights into novel nucleolar functions and processes. Investigating the interactions between plant viruses and the nucleolus will facilitate the design of novel strategies to control plant virus infections.

ABBREVIATIONS AGO BiFC CaMV CB CBL CCCVd CMV CP CP-RT DCL DEAD box protein DFC DRB FC GRV GAR GFP GC HC-Pro HIV IFN IBV IRES MFSV

argonaute protein bimolecular fluorescence complementation cauliflower mosaic virus Cajal body Cajal body-like structure coconut cadang cadang viroid cucumber mosaic virus coat protein coat protein-read through portion Dicer-like ribonuclease Asp-Glu-Ala-Asp family protein dense fibrillar component dsRNA-binding protein fibrillar centers groundnut rosette virus glycine and arginine rich domain green fluorescent protein granular component helper-component proteinase Human immunodeficiency virus interferon infectious bronchitis virus internal ribosome entry site maize fine streak virus

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MP miRNA NIa (b) NES NLS NMD NoLS NS1 ORF PABP PD PLRV PMTV pre-rRNA PSLV RDRP PSTVd PVA RNP rRNA sDMAs SMN snRNA siRNA snoRNA SPMV SRP TEV TGB ToLCJAV TuMV TYLCV VIRP1 VPg

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movement protein microRNA nuclear inclusion protein a (b) nuclear export signal nuclear localization signal nonsense-mediated decay nucleolar localization signal nonstructural protein 1 open reading frame poly(A) binding protein plasmodesmata potato leafroll virus potato mop top virus precursor rRNAs poa semilatent virus RNA-directed RNA polymerase potato spindle tuber viroid potato virus A ribonucleoprotein ribosomal RNA symmetric di-methyl arginines survival motor neuron gene product small nuclear RNA small interfering RNA small nucleolar RNA satellite panicum mosaic virus signal recognition particle tobacco etch virus triple gene block tomato leaf curl Java virus associated with DNA b satellite turnip mosaic virus tomato yellow leaf curl virus viroid RNA binding protein 1 viral genome-linked protein

I. INTRODUCTION The interactions between a virus and its host cell play a central role in the viral infection cycle. The analysis of virus–host interactions is critical for understanding the mechanisms of viral infections and for the development of novel antiviral strategies. Viruses are obligate intracellular pathogens with small genomes and, therefore, are reliant on subverting of the cellular

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functions and machineries to facilitate their own replication. The cell nucleus is one of the key features of eukaryotic cells. It is a highly dynamic, membrane-bound organelle that hosts major cellular events, including DNA replication, messenger RNA synthesis and processing, and ribosome subunit biogenesis (Trinkle-Mulcahy and Lamond, 2007). Given the pivotal role of the nucleus in cell host function, it is not surprising that viruses interact with this organelle and its compartments, and that such interactions play a crucial role in the virus infection cycle. Indeed, certain viruses including plant DNA containing begomoviruses (Rojas et al., 2001; Sharma and Ikegami, 2009), RNA reverse-transcribing caulimoviruses (CaMVs) (Haas et al., 2005), and negative-strand RNA rhabdoviruses belonging to the genus Nucleorhabdovirus (Tsai et al., 2005) replicate in the nucleus, and therefore it makes sense that these viruses modify nuclear structures to usurp some of the nuclear functions and provide an appropriate environment for their own replication. In contrast, single-stranded positive-sense RNA [(þ)ssRNA] viruses replicate in the cytoplasm. Therefore, the rational for RNA viruses targeting the nucleus and its compartments is not so evident. However, an increasing number of reports clearly show that cytoplasmic RNA viruses including plant viruses can also target nuclear compartments and, nucleoli, in particular (reviewed by Hiscox 2002, 2007; Greco, 2009). Thus, investigating why and how plant RNA viruses interact with the nucleolus to meet their own needs will contribute to the better understanding of molecular biology of plant viruses and facilitate the design of novel strategies for virus control. These studies may also teach us about the fundamental principles of plant nucleolar structure and functions. The nucleolus is a prominent subnuclear compartment formed around the clusters of genes (rDNA) coding for ribosomal RNA (rRNA), and is the site of rDNA transcription, rRNA processing, and ribosome assembly (Olson, 2004; Rubbi and Milner, 2003; Sirri et al., 2008). Indeed, for many years the exclusive function of the nucleolus was thought to be rRNA synthesis and ribosome biogenesis. However, several insights from the past decade have dramatically changed this traditional view and implicated the nucleolus in many other aspects of cell function such as cellcycle regulation, gene silencing, telomerase activity, senescence, stress responses, and biogenesis of multiple ribonucleoprotein (RNP) particles (Boisvert et al., 2007; Olson and Dundr, 2005; Sirri et al., 2008). This chapter briefly reviews the structure and composition of the nucleolus and, subsequently, data implicating the nucleolus in the infection cycles of animal and plant viruses and also viroids. Most of the current knowledge on nucleolar localization and functions of viral proteins has been gained in studies on animal viruses and has also been reviewed comprehensively (Greco, 2009; Hiscox, 2002, 2007). Therefore, the main emphasis of this review will be on what is known about the

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different aspects of interactions between plant virus proteins and the nucleolus, of which the functional significance in control of virus movement and interference with host antiviral defense has started to appear only recently. We also discuss recent findings on the potential role of Cajal bodies (CBs), another type of small subnuclear bodies structurally and functionally associated with the nucleolus.

II. STRUCTURE AND FUNCTIONS OF THE NUCLEOLUS The nucleus is a highly structured organelle responsible for chromosome organization, replication and division, for gene expression and regulation, and the integration of a vast array of activities required for cellular function. The nucleus contains chromatin-rich regions, made up of condensed heterochromatin, more dispersed euchromatin, and interchromatin regions, and chromatin organization is responsible for regulating gene expression, DNA replication, and chromosome segregation (Schneider and Grosscheld, 2007; Trinkle-Mulcahy and Lamond, 2008). In addition, it contains many other structures or bodies of different numbers and sizes which vary between cell types, at different stages in the cell cycle and under different conditions. The most prominent of these structures is the nucleolus but many other bodies [e.g., CBs, splicing speckles, paraspeckles, PML (pro-myeloid leukemia) bodies, etc.] have now been identified and are being characterized on the basis of their protein and RNA components and functions (Boisvert et al., 2007; Cioce and Lamond, 2005; Lamond and Spector, 2003; Matera et al., 2009; Rippe, 2007). The nucleolus is where rRNA genes are transcribed and processed, and rRNAs are assembled, along with ribosomal proteins, into the small and large subunits of the ribosome (Boisvert et al., 2007; Fatica and Tollervey, 2002; Granneman and Baserga, 2004). The nucleolus contains three different regions on the basis of their appearance in the transmission electron microscope: fibrillar centers (FC), the dense fibrillar component (DFC), and the granular component (GC). In the mammalian nucleolus, the FCs are small, lightly staining structures surrounded by the more densely stained DFC. The DFC in turn is surrounded a particulate region—the GC. The plant nucleolus tends to be more regular in its structure (usually near to spherical) than animal nucleoli. The organization of the nucleolar regions is different in plant nucleoli in that the DFC is not as densely stained as in animal nucleoli and occupies much more of the nucleolar volume (up to 70%). Transcription of precursor rRNAs (pre-rRNAs) occurs at multiple sites (200–400) within the DFC regions and pre-rRNAs undergo processing in the DFC and GC. Indeed, the localization of pre-rRNAs and different small nucleolar RNAs (snoRNAs) and proteins has allowed early and late pre-rRNA cleavage events to be

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correlated to the DFC and GC, respectively, suggesting a vectorial model for the production and maturation of rRNAs. Plant nucleoli also often contain a prominent central region called the nucleolar cavity (Brown and Shaw, 1998; Shaw and Brown, 2004). The function of the nucleolar cavity is currently unknown, and it remains to be seen whether the differences in organization of plant and animal nucleoli influence the type and range of other functions which nucleoli carry out and interactions with different viruses. The major function of the nucleolus is the transcription and processing of precursor rRNAs and ribosomal subunit assembly. The dynamic assembly pathway involves a series of intermediate pre-ribosomal complexes of the 40S and 60S ribosomal subunits and in addition to ribosomal proteins, requires up to 200 accessory proteins (Grannemann and Baserga, 2004). Processing of the pre-rRNAs involves both cleavage of the precursor transcript and nucleotide modifications, the majority of which are 20 -O-ribose methylation and pseudouridylation. Some cleavage reactions and the modifications require snoRNPs. SnoRNPs consist of a snoRNA and associated proteins (Kiss, 2002). By the nature of the major function in rRNA production and ribosomal subunit assembly, there is a huge flux of proteins and RNA complexes into and out of the nucleolus. The highly dynamic movement of proteins and complexes has been illustrated in animal cells by quantitative nucleolar proteomic analyses (Andersen et al., 2005; Lam et al., 2007). In particular, ribosomal proteins are highly expressed and rapidly accumulate in the nucleolus where they are incorporated into ribosomal subunits or rapidly degraded (Lam et al., 2007). Three of the most abundant and well-studied nucleolar proteins are fibrillarin, nucleolin, and B23. Fibrillarin, one of the major proteins of the nucleolus, is a core component of box C/D snoRNPs and is required for rRNA processing (Venema and Tollervey, 1999). Fibrillarin has methyltransferase activity directing 20 -O-ribose methylation of rRNA (Barneche et al., 2000; Cioce and Lamond, 2005; Matera and Shpargel, 2006). Fibrillarin is highly conserved in sequence, structure, and function in eukaryotes. The N-terminal region of fibrillarin comprises a glycine- and arginine-rich (GAR) domain (Barneche et al., 2000). The GAR domain is methylated at arginine residues (Liu and Dreyfuss, 1995) and is responsible for interactions with various proteins including the survival motor neuron (SMN) gene product ( Jones et al., 2001) and the nuclear DEAD (Asp-Glu-Ala-Asp) box protein p68 (Nicol et al., 2000). Fibrillarin contains a centrally located RNA-binding domain which together with the C-terminal helix domain and the intervening spacer constitutes a methyltransferase-like domain that contains an S-adenosyl methionine binding motif and is responsible for fibrillarin methyltransferase activity (Wang et al., 2000a). The C-terminal helix domain appears to target fibrillarin to CBs (Snaar et al., 2000). Although it is well established that fibrillarin plays a role in ribosome biogenesis within the nucleolus, its role in CBs is not

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well understood but it is presumably responsible for the 20 -O-ribose methylation of small nuclear RNAs (snRNAs). Nucleolin is an abundant, ubiquitously expressed protein, which is highly phosphorylated, methylated, and also can be ADP-ribosylated (Ginisty et al., 1999). Nucleolin is found in various cell compartments, and it is especially abundant in the nucleolus. Nucleolin has three well-defined domains. The N-terminal domain with alternating acidic and basic stretches is involved in rDNA transcription by interacting with rDNA repeats and histone H1 and in nuclear localization. The central portion is the RNAbinding domain, whereas the C-terminal part contains a GAR domain involved in interaction with the ribosomal proteins (Tuteja and Tuteja, 1998). Nucleolin is involved in ribosome biogenesis (Mongelard and Bouvet, 2007), affects transcription, processing and modification of rRNA and nuclear-cytosolic transport of ribosomal proteins and ribosomal subunits by shuttling between the nucleus and the cytoplasm (Tuteja and Tuteja, 1998). B23 is an abundant, multifunctional nucleolar phosphoprotein whose activities are proposed to play a role in ribosome assembly, binding to other nucleolar proteins, nucleocytoplasmic shuttling (Li et al., 1996), and possibly regulating transcription of rDNA by mediating structural changes in chromatin (Okuwaki et al., 2001). Two isoforms of B23 have been identified: the major form (B23.1) is predominately located in the nucleolus and the minor form (B23.2) resides in the cytoplasm (reviewed by Hiscox, 2002). Small proteins of less than 40–60 kDa can enter the nucleus through nuclear pore complexes by passive diffusion (Hiscox, 2007; Nigg, 1997). RNA-binding proteins that diffuse into the nucleus may therefore nonspecifically target the nucleolus as it contains a large amount of rRNA. The nuclear import of larger proteins is mediated by nuclear localization signals (NLSs), composed of one (monopartite) or two (bipartite) stretches of basic amino acid (arginine and/or lysine) residues of a given size (Hiscox, 2007; Macara, 2001). Nucleocytoplasmic shuttling proteins also contain specific nuclear export signals (NESs), usually leucine-rich amino acid motifs (Hiscox, 2007; Macara, 2001). How proteins may be further delivered to the nucleolus is poorly understood. The nucleolus does not have apparent membrane or other barriers, and entry into it does not require energy, unlike entry to the nucleus. It seems conceivable that viral proteins localize to the nucleolus, firstly, as a result of targeting the nucleus via classical NLS followed by direct or indirect interactions between the viral molecules (via various nucleolar localization signals, NoLSs) and components that make up the nucleolus (Hiscox, 2002, 2007). The structure of NoLSs is not well defined and depends on different factors including whether the protein associates with another nucleolar-bound protein or alternatively traffics to the nucleolus on its own

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or associates with RNA transcripts that are being transcribed in the nucleolus. At least in some cases, NoLS are rich in arginine and lysine residues and can overlap with NLSs (Hiscox, 2002, 2007). The second most studied nuclear body is CBs which are frequently associated with the nucleolus and found in animal and plant nuclei. CBs are involved in snRNP and snoRNP maturation and transport, with snRNPs and snoRNPs accumulating in CBs before appearing in speckles or the nucleolus, respectively (Cioce and Lamond, 2005). As mentioned above, spliceosomal snRNAs are also modified and contain 20 -O-ribose methylations and pseudouridines. Modification of nucleotides in snRNAs is guided by small CB-specific RNAs (scaRNAs) (Darzacq et al., 2002; Jady et al., 2003). A major component of CBs is the protein, coilin, which is required for their formation. In plants, mutants are available which knock out CBs or alter their size and number, two of which are due to mutations in coilin (Collier et al., 2006). The number of CBs per nucleus can vary in different cell types and is under developmental control (Cioce and Lamond, 2005). Besides rRNA transcription and processing and the production of ribosomal subunits, the nucleolus is also involved in many other aspects of RNA processing and RNP assembly as well as cellular functions (Boisvert et al., 2007; Olson, 2004; Pedersen, 1998; Rasˇka et al., 2006; Rubbi and Milner, 2003). For example, the nucleolus has a role in the maturation, assembly, and export of RNP particles. The signal recognition particle (SRP) has a nucleolar phase in its assembly with particular protein constituents of the SRP being localized to the nucleolus. Similarly, telomerase RNP, required for chromosome replication, may be assembled in the nucleolus, and the nucleolus may also have a role in sequestering telomerase RNP to avoid inappropriate nucleation of telomere structures. Spliceosomal snRNPs may also undergo part of their assembly in the nucleolus, and processing of pre-tRNAs and U6snRNA occurs in the nucleolus. In addition, the nucleolus is a site of sequestration of particular proteins to regulate, for example, the cell cycle or cell death, and acts as a sensor of cellular stress (Boisvert et al., 2007; Olson, 2004; Pedersen, 1998; Rasˇka et al., 2006; Rubbi and Milner, 2003). Characterization of the protein composition of the nucleolus under different conditions has provided support for or suggested new functions for the nucleolus. Initial proteomic analyses of human cells identified a few hundred proteins which included not only well-known nucleolar proteins such as ribosomal proteins, fibrillarin, nucleolin, B23, etc. but also, for example, splicing and translation factors (Andersen et al., 2002). Advances in proteomics has now allowed the identification of around 4500 proteins in the human nucleolar proteome (Ahmad et al., 2008) and the dynamic behavior of nucleolar components and complexes can now be determined (Andersen et al., 2005; Lam et al., 2007). In plants, a partial

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proteomic analysis of the nucleolus identified many expected ribosomal and nucleolar proteins but also found splicing and translation factors (Pendle et al., 2005). In particular, exon junction complex proteins (associated with mRNAs following splicing) were identified in the nucleolar proteome and in the nucleolus by fluorescence microscopy (Pendle et al., 2005). One of these proteins, eIF4A-III, a core protein of the exon junction complex, was shown to redistribute from the nucleoplasm to the nucleolus and finally to splicing speckles under stress conditions of hypoxia (Koroleva et al., 2009). Similar dynamic distribution, involving the nucleolus, of proteins that interact with mRNAs has been demonstrated and the nucleolus and CBs have been shown to be involved in U1snRNP production in plants (Lorkovic´ and Barta, 2008; Tillemans et al., 2006). In plants, novel functions for the plant nucleolus, CBs, and another largely nucleolar-associated body, the D-body, have been described. Firstly, the production of heterochromatic siRNAs, which are involved in transcriptional silencing, is thought to occur in a region of the nucleolus or in D bodies due to the localization of protein components of the machinery and of siRNAs (Pontes and Pikaard, 2008). Secondly, maturation of microRNAs (miRNAs) may occur in D-bodies as precursor miRNAs as well as Dicer-like ribonuclease 1 (DCL1) have been located to these structures (Pontes and Pikaard, 2008). Similarly, in mammalian cells, some precursor and mature miRNAs have recently been found in the nucleus with some being enriched in the nucleolus (Politz et al., 2009; Scott et al., 2009). Of particular interest was the enrichment of some precursors in the GC suggesting that miRNA processing could occur in the nucleolus, or these miRNAs may be involved in ribosome synthesis or other nucleolar functions (Politz et al., 2009). The link between the nucleolus and miRNA production is illustrated by the evolutionary relationship between some snoRNAs and miRNA precursors. For example, a human snoRNA was shown to be processed by Dicer to generate small RNAs which were associated with argonaute proteins (AGOs) and caused reduced expression of gene targets (Ender et al., 2008). In addition, numerous snoRNA-derived small RNAs from different organisms (including Arabidopsis) were associated with components of RNA silencing pathways (Taft et al., 2009) and many miRNA precursors have retained snoRNA features (Scott et al., 2009). Finally, a third novel function for the plant nucleolus is in mRNA biogenesis, surveillance, or nonsensemediated decay (NMD). Biochemical fractionation of nucleoplasm and nucleoli and subsequent sequencing of isolated cDNAs have shown that the plant nucleolus not only contains mRNAs but that aberrant mRNAs are enriched in the nucleolus (Kim et al., 2009). The aberrant mRNAs show splicing defects, the majority of which would introduce premature termination codons and therefore be expected to be substrates for NMD. Using upf mutants (affected UPF proteins, key components of the NMD

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mechanism), the correlation between enrichment of aberrant mRNAs in the nucleolus and turnover by NMD was corroborated and was further supported by the localization of the NMD proteins, UPF2 and UPF3, to the nucleolus (Kim et al., 2009). Before the observation that the plant nucleolus contained mRNAs and aberrant mRNAs, some spliced mRNAs (e.g., c-myc) were localized to the nucleolus while their unspliced versions were found in the nucleoplasm in mammalian cells (Pedersen, 1998; Olson, 2004). The nucleolus has also been associated with mRNA export on the basis of accumulation of polyAþ RNA upon disruption of export factors, nucleolar structure, or stress conditions in yeast and animal cells, although this could reflect increased polyadenylation prior to degradation (Ideue et al., 2004; Pederson, 1998; Schneiter et al., 1995). More recently, the nucleolus has been proposed to be involved in the formation of mRNPs which are localized to specific regions of the cytoplasm for translation. In yeast, ASH1 mRNA enters the nucleolus bound to specific RNA-binding proteins at which time translation repressor proteins are loaded onto the mRNA. The mRNP is then exported to the cytoplasm, transported to its final destination, and then translation is activated (Du et al., 2008; Jellbauer and Jansen, 2008). A similar trafficking pathway may also operate in mammals for mRNPs associated with the nucleolar protein, Staufen, which is involved in transport of mRNAs in neurons ( Jellbauer and Jansen, 2008). Thus, the nucleolus has numerous functions related to RNA biogenesis and different RNA processing and RNP maturation pathways. It contains many RNA-interacting proteins involved in these processes, including highly abundant RNA-binding proteins (such as fibrillarin and nucleolin) involved in rRNA and ribosomal subunit production. It is therefore a dynamic hub of RNA processing activity, RNA:protein interaction and complex formation. These characteristics, in addition to the potential involvement of the nucleolus in mRNA biogenesis and, particularly, the transport of mRNAs and mRNPs to and from the nucleolus to other parts of the cell, make the nucleolus a prime target for exploitation by viruses. It is therefore not surprising that viruses have taken advantage of the nucleolus and nucleolar components for production and distribution of viral RNAs and RNPs.

III. WHAT WE HAVE LEARNED ABOUT INTERACTIONS OF VIRUSES WITH THE NUCLEOLUS FROM ANIMAL VIROLOGY Several excellent reviews have documented various aspects of the involvement of the nucleolus in the infection cycle of animal and human viruses (Greco, 2009; Hiscox, 2002, 2007; Matthews and Olson, 2006; Stark

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and Taliansky, 2009). This section therefore briefly summarizes the main findings in this research area to illuminate the nucleolar functions involved in virus infections that are conserved between plant and animal kingdoms.

A. Viruses that replicate in the nucleus Certain animal viruses such as DNA containing adenoviruses that replicate in the nucleus interact with and disrupt the nucleolus. As a result, synthesis of rRNA is disrupted in adenovirus-infected cells. Adenovirus infection also causes the redistribution of nucleolin and B23 (Matthews, 2001). Interestingly, B23 has been shown to stimulate adenovirus replication. RNA virus replication may also be facilitated by nucleolar proteins. Indeed, the HDAg protein encoded by hepatitis delta virus (a ()ssRNA virus which is a subviral satellite of the DNA virus, hepatitis B virus) also binds to nucleolin and B23. It has been proposed that HDAg interacts with both these proteins in a complex that promotes viral RNA replication, presumably as a result of the helicase activity of nucleolin (Huang et al., 2001). Nucleolar proteins may also be involved in virus assembly and egress. For example, accumulation and assembly of structural (Cap) proteins of adeno-associated virus take place in nucleoli (Bevington et al., 2007). Further export of the assembled Cap proteins from the nucleolus is mediated by another type of virus-encoded packaging (Rep) proteins, which should form a complex with Cap proteins. Remarkably, formation of this complex is mediated by B23. After export of complexes containing Rep, B23, and Cap proteins from the nucleolus to the nucleoplasm, viral DNA encapsidation occurs (Bevington et al., 2007). Another animal virus, the ()ssRNA containing Borna disease virus, uses the nucleolus as a site of genome replication (Pyper et al., 1998). An RNA-binding protein encoded by this virus has the appropriate trafficking signals for import and export to and from the nucleus (Cros and Palese, 2003)

B. Viruses replicating in cytoplasm Many proteins encoded by animal viruses that replicate mainly or exclusively in the cytoplasm have also been shown to localize to the nucleolus, cause the relocalization of nucleolar proteins and disruption of nucleolar architecture and function. These include the (þ)ssRNA virus proteins such as nucleocapsid proteins encoded by coronavirus and arterivirus, the dengue virus core protein, the alphavirus capsid protein and nonstructural nsP2 protein, the ()ssRNA virus proteins, including the influenza virus nucleoprotein (NP) and nonstructural protein 1 (NS1),

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Newcastle disease virus matrix protein, and retrovirus proteins such as human immunodeficiency virus (HIV) Rev and the transactivator Tat proteins (reviewed by Greco, 2009; Hiscox, 2002, 2007). Although functional relevance of why these proteins localize to the nucleus or nucleolus and how this relates to their functions in virus replication in many cases is largely unknown, several reports reveal significant progress in this area as exemplified below. Infection of cells with poliovirus (picornavirus) results in the inactivation of the nucleolar protein, RNA polymerase I upstream binding (transcription) factor, which inhibits transcription of rRNAs in the host cell (Banerjee et al., 2005). Poliovirus is also able to interact with nucleolin causing its selective redistribution from the nucleolus to the cytoplasm (Waggoner and Sarnow, 1998). After relocalization, nucleolin binds to the internal ribosome entry sites (IRESs) at the 50 untranslated region of poliovirus genomic RNA, and this interaction stimulates IRES-dependent translation (Hellen and Sarnow, 2001). This also occurs in hepatitis C virus (Izumi et al., 2001) and represents an alternative translation initiation strategy as compared to the classical eukaryotic Cap-dependent translation (Hellen and Sarnow, 2001). Nucleolin is also able to interact with the 30 -untranslated region of poliovirus RNA, which controls synthesis of negative strand RNA (Waggoner and Sarnow, 1998). Interaction of picornaviruses with the nucleolus could also down-regulate host gene expression. For example, the human rhinovirus 3C protease precursors that are localized in the nucleolus at early stages of the infection inhibit cellular RNA by cleavage of vital transcription factors (Amineva et al., 2004). The nucleocapsid proteins encoded by porcine arterivirus and avian coronavirus (infectious bronchitis virus, IBV) interact with nucleolin and fibrillarin (Chen et al., 2002; Yoo et al., 2003), and as a result may disrupt the normal functions of these proteins. For instance, by altering the distribution of fibrillarin, viruses might be reducing polI transcription, that is, the synthesis of rRNA, as blocking fibrillarin with antibody prevented its translocation to nucleoli and resulted in the reduction or inhibition of polI transcription (Fomproix et al., 1998). The IBV infection leads to disruption of the nucleolus (Dove et al., 2006a) and arrest of the cell cycle and cytokinesis (Chen et al., 2002; Dove et al., 2006b; Wurm et al., 2001). Therefore, disruption of nucleolar architecture and function might be common in cells infected with viruses interacting with the nucleolus. The loss of essential nucleolar function in its turn may play an important role during virus infection toward an active production of the virus. One of the most studied viruses in terms of viral interactions with the nucleolus is HIV, a retrovirus. HIV RNAs are reverse transcribed in the cytoplasm of infected cells and trafficked to the nucleus. After transcription in the nucleus, progeny RNA molecules are transported back to the

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cytoplasm. One of the functions of the Rev protein is to export unspliced or partially spliced viral mRNA from the nucleus (reviewed by Greco, 2009; Hiscox, 2007). Before nuclear export, HIV RNA passes through the nucleolus. Rev binds to a cis-acting RNA element (Rev-response element), which is found in all unspliced and incompletely spliced viral mRNAs, and this promotes the translocation of these RNAs from the nucleus (Canto´-Nogue´s et al., 2001; Michienzi et al., 2000). The nucleolus plays a central role in this process, and the nucleolar trafficking of Rev and viral RNA is critical for the outcome of infection. Thus, many animal viruses, whether they replicate or not in the nucleus, have evolved a nucleolar phase for part of their infection cycle to prevent unwanted interference from the cell. Alternatively, they use nucleolar functions for their own benefit. Recruitment of nucleolar proteins is especially beneficial for viruses, and in particular for RNA containing viruses, as these proteins possess many crucial functions in cellular RNA biosynthesis, processing, translation, and trafficking. Indeed, during virus infections of mammalian cells, various viral components traffic to and from the nucleolus where they interact with different host factors. Certain nucleolar proteins are redistributed into other cell compartments or are modified, and some cellular proteins are relocalized in the nucleolus of infected cells (reviewed by Greco, 2009; Hiscox, 2002, 2007). Well-documented studies have established that several of these nucleolar modifications play a role in some steps of the viral infection cycle such as viral attachment and entry, intracellular trafficking, transcription, translation, replication, virus assembly, and regress (reviewed by Greco, 2009). The virally induced nucleolar modifications could also affect fundamental cellular pathways including the initiation of transcription from the DNA promoter of the rRNA genes, cell-cycle regulation, and apoptosis (reviewed by Greco, 2009). Although some steps (replication, translation) of the infection cycles of plant and animal/human viruses are essentially similar, there is a fundamental difference in some other mechanisms employed to enter host cells and spread from cell to cell between viruses infecting animal cells and viruses infecting plants. This is because animal cells are separated by barriers far less formidable than the thick, rigid, and impermeable cell walls consisting of cellulose and pectin that separate plant cells from one another. Other differences relate to defense strategies employed by humans/animals and plants against viral infections. For example, in mammals the interferon (IFN) pathway plays a key role in the innate antiviral immune response, whereas plants do not display such an activity. Instead, plants primarily rely on other natural defense strategies such as RNA silencing. Interestingly, functional links between plant virus infection cycle and the nucleolus have been described for both common and plant-specific virus infection steps.

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IV. NUCLEOLAR FUNCTIONS OF PLANT VIRUS PROTEINS AND VIROIDS Certain plant virus proteins localize to the nucleolus with examples from single-stranded DNA viruses, para-retroviruses and negative-strand and positive-strand RNA viruses (Table I). The most common technique for studying the nucleolar targeting of plant virus proteins is based on the confocal microscopy localization of the proteins which have been tagged with a fluorescent fusion protein (such as green fluorescent protein, GFP). Such proteins are usually larger than the size-exclusion limit ( 40– 60 kDa) and hence prevented from nonspecific protein diffusion into the nucleus. Moreover, in many cases the specific nucleolar localization of plant virus proteins has been supported by identification of NoLSs (Table II). Although no overall conserved nucleolar trafficking motif has been identified in these NoLSs, they presumably resemble host NoLSs. Thus plant viral nucleolar trafficking might use a form of molecular mimicry as has earlier been proposed for animal viruses (Rowland and Yoo, 2003). Like host NoLSs, plant virus NoLSs may contain single [ToLCJAV CP, PLRV CP (CP-RT), CMV 2b] or bipartite [GRV ORF3, PVA NIa (VPg)] motifs which are usually characterized by basic amino acid stretches (Table II). However, in the case of GRV ORF3, in addition to the arginine-rich domain (NLS), a leucine residue at position 149 (L149) residing inside the leucine-rich domain (NES) is also essential for nucleolar targeting of ORF3 protein (Table II; Kim et al., 2007a). The fact that viral proteins contain NoLSs is a strong indication that viruses have evolved specific nucleolar functions. Viral proteins might also traffic to the nucleolus through association with host proteins. One example of such an association may be the interaction of various plant virus proteins with the major nucleolar protein, fibrillarin. The first description of this interaction was the demonstration that the umbravirus GRV ORF3 long-distance movement protein (MP) binds to fibrillarin in vivo and in vitro (Kim et al., 2007a,b). For example, the leucine-rich domain (and L149, in particular) of ORF3 is involved in direct interaction with fibrillarin (Kim et al., 2007b). Mutations in the leucine-rich domain prevent fibrillarin from binding to ORF3 and nucleolar trafficking (Kim et al., 2007b). By implication, this may relate fibrillarin binding to nucleolar trafficking. Other plant virus MPs which interact with fibrillarin are the pomovirus PMTV triple gene block protein 1 (TGB1) and hordeivirus PSLV TGB1 (N. O. Kalinina and D. Rakitina, unpublished results). As their name suggests, MPs are involved in virus spread in infected plants, and the potential role of fibrillarin in this process will be discussed in Section IV.B. The multifunctional PVA (potato virus A)-encoded viral genomelinked protein (VPg) is also able to interact with fibrillarin (Rajama¨ki and

TABLE I Examples of plant virus proteins and viroids that localize to the nucleolus Genus

Virusa

DNA (single-stranded) viruses Begomovirus TYLCV ToLCJAV

a

b

Protein/RNAb

Reference (s)

CP CP

Rojas et al. (2001) Sharma and Ikegami (2009)

RNA reverse transcribing virus (para-retrovirus super-group) Caulimovirus CaMV P6

Haas et al. (2005)

Negative-stranded RNA virus Nucleorhabdovirus MFSV

N and P proteins

Tsai et al. (2005)

Positive-stranded RNA viruses Potyvirus TEV

NIa (VPg); NIb; P3

Restrepo et al. (1990), Baunoch et al. (1991), Langenberg and Zhang (1997) Beauchemin et al. (2007) Rajama¨ki and Valkonen (2009) Ryabov et al. (1998), Ryabov et al. (2004) Haupt et al. (2005) Qi et al. (2008) Gonza´lez et al. (2010), Mackenzie and Tremaine (1988) Torrance (personal communication) NOK (unpublished results)

Umbravirus Polerovirus Satellite virus (Panicovirus) Cucumovirus

TuMV PVA GRV PLRV SPMV CMV

NIa (VPg) NIa (VPg) ORF3 CP (CP-RT) CP 2b, 3a (MP)

Pomovirus Hordeivirus

PMTV PSLV

TGB1 TGB1

Viroid Pospiviroid

PSTVd, CCCVd

RNA

Schumacher et al. (1983), Harders et al. (1989), Bonfiglioli et al. (1996), Qi and Ding (2003)

Virus acronyms: TYLCV, tomato yellow leaf curl virus; ToLCJAV, tomato leaf curl Java virus associated with DNA b satellite; CaMV, cauliflower mosaic virus; MFSV, maize fine streak virus; TEV, tobacco etch virus; TuMV turnip mosaic virus; PVA, potato virus A; GRV, groundnut rosette virus; PLRV, potato leafroll virus; SPMV, satellite panicum mosaic virus; CMV, cucumber mosaic virus; PMTV, potato mop top virus; PSLV, poa semilatent virus; PSTVd, potato spindle tuber viroid; CCCVd, coconut cadang cadang viroid. CP, coat protein; P6, CaMV multifunctional protein; N protein, nucleocapsid protein; P protein, phosphoprotein; NIa (b), nuclear inclusion protein a (b); VPg, viral genomelinked protein; P3, potyviral unstructural protein with no well-characterized function; ORF3, open reading frame 3 protein; CP-RT, coat protein-read through portion; 2b, CMV silencing suppressor; MP, movement protein (3a, CMV MP); TGB1, triple gene block protein 1.

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TABLE II Examples of plant virus proteins that contain nucleolar localization signals (NoLS) Protein

ToLCJAV CP PVA NIa (VPg) GRV ORF3 PLRV CP (CP-RT) CMV 2b

Amino acid position

NoLS

Reference (s) a

16–20

KVRRR (NLS)

Sharma and Ikegami (2009) Rajama¨ki and Valkonen (2009)

4–9 41–50 108–122 148–156 17–31

KRQRQK (NLS I)b KKGKTKGKTH (NLS II)b RPRRRAGRSGGMDPR Ryabov et al. (2004) Kim et al. (2007a) LLPSLLNTL (NES)c PRRRRRQSLRRRANR Haupt et al. (2005)

22–27

KKQRRR (NLS1)a

Gonza´lez et al. (2010)

Virus acronyms and other abbreviations are as in Table I. a NLSs are also required for the nucleolar localization. b Nuclear and nucleolar localization is controlled independently by the same NLS regions. c L149 (italic) residing inside the ORF3 NES is also essential for nucleolar localization of the ORF3 protein.

Valkonen, 2009). However, the role of this interaction is likely to be different from that in virus movement as this process is not compromised by fibrillarin depletion (Rajama¨ki and Valkonen, 2009; Section IV.C). Another example of a viral protein that interacts with fibrillarin in the nucleolus is a silencing suppressor of CMV (cucumber mosaic virus), the 2b protein (Gonza´lez et al., 2010; Section IV.C). Collectively, these data indicate that interaction with fibrillarin is a general property of various plant virus proteins that is not restricted to one or two virus taxonomic groups. However, such interactions may have quite diverse molecular implications for different viruses being required at various phases of virus infection cycle. This may also suggest novel, unexpected natural functions for fibrillarin that are hijacked or affected by plant viruses at different stages of infection for needs of the viruses.

A. Nucleolus in replication of viroids and viruses Viroids are small, circular, self-replicating, and non-coding RNA molecules that cause plant diseases (reviewed by Tabler and Tsagris, 2004). Viroids do not replicate in the cytoplasm like conventional plant RNA viruses but replicate in either the nucleus or the chloroplast. Nuclear viroids such as PSTVd and CCCVd have a nucleolar phase in their replication cycle (Table I; Qi and Ding, 2003; Schumacher et al., 1983).

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In early experiments using in situ hybridization, PSTVd RNA [including both (þ) and () RNA strands] was localized in nucleoli of nuclei isolated from infected plants (Harders et al., 1989), suggesting that the nucleolus is the replication site of the viroid. However, later using improved sample preparation and in situ hybridization protocols, Qi and Ding (2003) have found that the () strand of PSTVd localizes in the nucleoplasm but not in the nucleolus. By contrast, the (þ) strand of PSTVd localizes in the nucleolus as well as in the nucleoplasm, with various distinct spatial patterns. These experiments are suggestive of successive stages of nuclear involvement in viroid replication: (1) synthesis of the () and (þ) strands of PSTVd occurs in the nucleoplasm; (2) the () strand RNA is anchored in the nucleoplasm; (3) the (þ) strand RNA is transported selectively into the nucleolus; and (4) some (þ) strand RNA traffics from the nucleolus back into the nucleoplasm and further into the cytoplasm for spreading into neighboring cells. The significance of (þ) strand PSTVd RNA trafficking into the nucleolus remains to be determined. However, nucleolar machineries for processing of rRNAs, snoRNAs, and tRNAs make the nucleolus an attractive candidate site for PSTVd processing. The specific intranuclear localization of () and (þ) strands of PSTVd may have some implication for pathogenicity. Assuming that the () and (þ) strands each can interact with specific cellular factors to disrupt normal cell functions to cause symptoms, the differential subnuclear localization of the (þ) and () strands of PSTVd may suggest different cellular targets for these RNAs. (þ) strand of PSTVd RNA has been shown to interact with a bromodomain-containing protein (a member of a family of transcriptional regulators associated with chromatin remodeling), termed viroid RNA binding protein 1 (VIRP1) (Martı´nez de Alba et al., 2003; Tabler and Tsagris, 2004). VIRP1 contains an NLS and therefore might transfer the viroid RNA to the nucleus and bring it into contact with transcription units associated with chromatin (Martı´nez de Alba et al., 2003; Tabler and Tsagris, 2004). However, mechanisms for selective nucleolar trafficking of (þ) strand PSTVd are still unknown. The nucleolus and its factors may be used not only by viruses and viroids for their own needs in replication, but also by plant host defense systems. For example, one of the major nucleolar proteins, nucleolin, binds to the 30 non-coding region of the tomato bushy stunt virus, tombusvirus, RNA ( Jiang et al., 2010). This leads to significant inhibition of tombusviral RNA replication and may thus represent one of the innate immunity systems of plant hosts.

B. The nucleolus and plant virus movement Plant viruses enter cells either through sites of mechanical injury to plant tissues or during the feeding by a specific vector organism (insect, nematode, or soil microbes belonging to protocists). To induce disease, after

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replication in the initially infected cells, plant viruses must spread to the rest of the plant. The systemic spread of plant viruses proceeds in two stages: (i) cell-to-cell movement through plasmodesmata (PD) and (ii) long-distance movement through vascular tissues. First, the virus (in the form of virions or nucleic acid protein complexes) moves intracellularly from the sites of replication to PD, which are unique intercellular membranous channels that span cell walls linking the cytoplasm of contiguous cells. The virus then transverses the PD to spread intercellularly. It is generally accepted that viral cell-to-cell movement involves virus-encoded MPs as well as host-encoded components (reviewed by Lucas, 2006). Virus systemic movement between organs (long-distance movement) occurs through the phloem, the specialized vascular system used by plants for the transport of assimilates and macromolecules (reviewed by Lucas, 2006; Oparka, 2004). Viruses enter, move through, and exit from the vascular system, which is usually surrounded by bundle sheath cells and contains various cell types including vascular parenchyma cells, companion cells, and enucleate sieve elements. Thus, transport of a virus into and within vascular tissue implies movement from mesophyll cells to bundle sheath cells, from bundle sheath cells to vascular parenchyma and/or companion cells, and into sieve elements. Virus exit from vascular tissue presumably involves the same steps in reverse order. Coat protein (CP) is essential for efficient long-distance transport of plant viruses, with only few exceptions (reviewed by Lucas, 2006).

1. Umbraviruses One of the most well-studied plant viruses in terms of viral interactions with the nucleolus is GRV, an umbravirus [(þ)ssRNA virus]. Umbraviruses differ from most other viruses in that they do not encode a CP such that conventional virus particles are not formed in infected plants. Umbraviral genomes encode at least three proteins. In GRV, two ORFs at the 50 -end of the RNA are expressed by a frameshift mechanism as a single protein that appears to be an RNA replicase (Taliansky and Robinson, 2003). The other ORFs (ORF3 and ORF4) overlap each other. ORF4 encodes the MP that mediates the cell-to-cell movement of viral RNA via PD (Ryabov et al., 1998). ORF3 protein is the long-distance movement factor that facilitates trafficking of viral RNA through the phloem (Ryabov et al., 1999). Umbraviral ORF3 proteins (26–29 kDa) show no significant similarity with any other recorded or predicted proteins (Taliansky and Robinson, 2003). The GRV ORF3 protein interacts with viral RNA in vivo to form filamentous RNP particles, which have elements of regular helical structure, but not the uniformity typical of virus particles (Taliansky et al., 2003). The RNPs accumulate in cytoplasmic inclusions which have been detected in all cell types and were

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abundant in phloem cells (Taliansky et al., 2003). They serve to protect viral RNA and move it through the phloem to cause systemic infection. Remarkably, in addition to its presence in the cytoplasm, the ORF3 protein was also found in nuclei and predominantly in nucleoli (Ryabov et al., 1998, 2004). Studies of the biology of GRV infection have provided molecular insights into how and why viruses may target the nucleolus (Canetta et al., 2008; Kim et al., 2007a,b). It has been demonstrated that the GRV ORF3 protein traffics to the nucleolus via a mechanism involving the reorganization of CBs into multiple Cajal body-like structures (CBL) and their fusion with the nucleolus. Nucleolar localization and further trafficking of ORF3 protein from the nucleolus back to the cytoplasm is essential for the umbravirus infection. The integral connection between nucleolar targeting of the ORF3 protein and its biological function in virus long-distance spread is demonstrated by mutagenesis of the arginine- and leucine-rich domains that block nucleolar localization or nuclear export of the ORF3 protein, and which prevent the formation of cytoplasmic viral RNPs and their long-distance movement (Kim et al., 2007a). Although the mechanisms by which the ORF3 protein targets CBs and produces CBLs are unknown, it could be suggested that targeting of CBs by the ORF3 protein may utilize elements of existing CB-trafficking pathways. For example, part of the maturation pathway of snRNPs in mammalian cells occurs in the cytoplasm and involves a complex containing the SMN protein (SMN complex) which together with the snRNP are reimported into the nucleus and targeted to CBs (Matera and Shpargel, 2006; Navascues et al., 2004; Sleeman and Lamond, 1999). Particular snRNP proteins contain modified symmetric di-methyl arginines (sDMAs) which enhance the formation of snRNPs and interaction with SMN (Paushkin et al., 2002). Preliminary experiments have identified sDMAs in the ORF3 protein (M. Taliansky, unpublished results), suggesting that targeting of the ORF3 protein to CBs could involve interactions with the SMN protein. Although SMN protein has yet to be identified in plants, the existence of an orthologue has been suggested (Collier et al., 2006). The formation of CBLs may involve either fragmentation of CBs into multiple bodies by the ORF3 protein or the redistribution of CB components into new structures containing the ORF3 protein. Interestingly, the multiple CBL phenotype, described here, is similar to that of the poly Cajal bodies (pcb) mutant of Arabidopsis (Collier et al., 2006). As the protein normally encoded by a gene controlling the pcb phenotype, appears to regulate CB formation, ORF3 protein may interfere with the function of this or other proteins to affect the integrity and number of CBs in nuclei. The second possibility is that the ORF3 protein causes the redistribution of CB components, such as coilin, U2B00 and fibrillarin, to form CBLs with the ORF3 protein.

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ORF3 protein trafficking to the nucleolus uses a novel pathway of nucleolar import by causing the fusion of CBLs with the nucleolus. The physical and functional association of the nucleolus and CBs is welldocumented and is controlled by complex molecular interactions among CB and nucleolar proteins such as coilin, SMN, fibrillarin, and Nopp140 (Cioce and Lamond, 2005; Ogg and Lamond, 2002). Expression of mutant versions of some of these proteins has profound effects on CB structure and function causing disruption or dispersal and compositional changes ( Jones et al., 2001; Pellizzoni et al., 2001; Tucker et al., 2001). Moreover, phosphorylation of coilin is an important factor determining physical interactions and trafficking of CBs (Cioce and Lamond, 2005; Ogg and Lamond, 2002). For example, CBs form within the nucleolus of HeLa cells upon treatment with okadaic acid (an inhibitor of protein phosphatase) and with transient expression of coilin mutated at a single serine residue (Lyon et al., 1997; Sleeman et al., 1998). CBs have also been observed within nucleoli in human breast carcinomas (Ochs et al., 1994) and in liver cells of hibernating dormice (Malatesta et al., 1994). Therefore, the ORF3 protein may interfere with normal protein–protein interactions or posttranslational modifications causing the reorganization and fusion of CBs with the nucleolus. The last stage of the nuclear voyage of the ORF3 protein is its nuclear export leading to the redistribution of some fibrillarin from the nucleolus to cytoplasm (fibrillarin normally does not accumulate in cytoplasm) (Kim et al., 2007a). This redistribution is mediated by the direct interaction between the ORF3 protein and fibrillarin (Kim et al., 2007b). Taking into account the long-distance movement function of GRV ORF3, it could be expected that fibrillarin is directly involved in this particular viral function. Further support of a role for fibrillarin in umbravirus systemic infection has been provided by the fibrillarin knock-down experiments (Kim et al., 2007b). Silencing of the fibrillarin gene has indeed prevented formation of RNP particles and long-distance movement of GRV but does not affect viral replication or cell-to-cell movement. Thus, it has been concluded that fibrillarin is needed for formation of cytoplasmic RNP particles that are capable of long-distance movement and causing systemic viral infection such that the redistribution of fibrillarin with ORF3 protein is a prerequisite to form RNPs (Kim et al., 2007a,b). Further experiments have shown that fibrillarin, in combination with ORF3 protein and viral RNA in vitro, produced filamentous RNP structures with structural properties similar to in vivo RNPs (as discussed in detail in Section IV.E). Fibrillarin, an RNA-binding protein, may bind the viral RNA or act as a chaperone to permit or catalyze the regular assembly of proteins around viral RNA. Although fibrillarin and ORF3 protein are sufficient to form functional viral RNP particles in vitro, additional proteins may also be associated with the in vivo particles. When formed in

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phloem companion cells, the viral RNPs are able to enter sieve elements and move through the plant to cause systemic infection. Hence formation of the fibrillarin-ORF3 protein complexes appears to be the key prerequisite for formation of GRV RNP particles and their long-distance movement through the phloem.

2. Poleroviruses Poleroviruses are (þ) ssRNA viruses that are mainly restricted to cells in the vascular system. Besides a major CP PLRV encodes a minor CP, an extended version of the major CP produced by occasional translational ‘‘readthrough’’ of the CP gene (CP-RT) (Bahner et al., 1990). Both CP and CP-RT contain the same NoLS, and are targeted to the nucleolus when they are individually expressed in plant tissues (Haupt et al., 2005). However, CP-RT but not PLRV CP loses its nucleolar localization in the presence of replicating PLRV. It has been suggested that the CP-RT protein does not accumulate in the nucleolus in the presence of PLRV infection because PLRV RNA or PLRV-encoded or -induced factors retain this protein outside the nucleolus (Haupt et al., 2005). Like GRV, PLRV has been unable to cause systemic infection in the fibrillarin-silenced plants, although accumulation of PLRV in the inoculated leaves was not affected (Kim et al., 2007b) suggesting that fibrillarin is also involved in PLRV long-distance movement.

3. Viruses that require a TGB for movement A role of nucleolar functions in systemic infection has also been suggested for plant viruses that require TGB for movement. The genomes of some viruses, such as potexviruses, hordeiviruses, and pomoviruses, contain so-called triple gene block, TGB that encodes three proteins required for cell-to-cell and long-distance movement (reviewed by Morozov and Solovyev, 2003). The TGB1 protein encoded by the TGB-containing virus, PMTV (pomovirus) localizes to the nucleus and nucleolus. Deletion of 84 Nterminal amino acids abrogates its nucleolar localization. Northern blots of RNA from inoculated and upper non-inoculated leaves of plants infected with clones carrying the TGB1 N-terminal deletion mutant reveal that long distance movement of viral RNAs has also been abolished, but this mutant is still competent for cell-to-cell movement (L. Torrance, personal communication). Moreover, the PMTV TGB1 protein has been shown to interact with fibrillarin in vitro. Interestingly, TGB1 protein encoded by the hordeivirus, PSLV can also interact with fibrillarin both in vitro and in vivo (N. O. Kalinina, unpublished results). Thus it appears that some TGB-encoding viruses such as PMTV and PSLV may represent another example of plant viruses that require association with the nucleolus to control long distance movement of their genomes.

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C. Nucleolar targeting for interference with host antiviral defense Potyviruses form the largest group of plant-infecting RNA viruses (Rajama¨ki et al., 2004). They have a polyadenylated (þ) ssRNA genome of ca. 9500–10,000 nucleotides that is encapsidated into a 680–900 nm long, filamentous particle. The genome is translated into a large polyprotein of about 3000–3350 amino acids, which is subsequently processed into up to ten mature proteins by three virus-encoded proteinases (Rajama¨ki et al., 2004). Additionally, a 25-kDa protein produced from the P3 cistron by frameshifting was recently identified (Chung et al., 2008). Potyviruses replicate in membranous structures in the cytoplasm (Cotton et al., 2009; Schaad et al., 1997a). However, two potyviral replication-associated proteins, the nuclear inclusion protein a (NIa) and nuclear inclusion protein b (NIb), accumulate in the nucleus of virus-infected cells (Baunoch et al., 1991; Dougherty and Hiebert, 1980; Hajimorad et al., 1996; Knuhtsen et al., 1974; Restrepo et al., 1990). In addition, NIa localizes in the nucleolus and CBs (Baunoch et al., 1991; Beauchemin et al., 2007; Rajama¨ki and Valkonen, 2009; Restrepo et al., 1990). The NIb of TEV (tobacco etch virus) has also been detected in the nucleolus (Baunoch et al., 1991; Restrepo et al., 1990). Potyviruses may also induce nuclear inclusions, which consist of NIa and NIb (Baunoch et al., 1991; Dougherty and Hiebert, 1980; Edwardson and Christie, 1983; Knuhtsen et al., 1974; Martin et al., 1992). The significance of these nuclear inclusions is unknown but they may simply represent inactivated protein complexes, because they seem to form only at late stages of virus infection (Baunoch et al., 1991). NIa is multifunctional and consists of two domains. The N-proximal half of NIa is the VPg that is covalently linked to the 50 -end of viral genomic RNA (Oruetxebarria et al., 2001; Siaw et al., 1985). The C-proximal part (NIa-Pro) is the main viral proteinase (Dougherty et al., 1989). The two domains are separated by a suboptimal proteolytic cleavage site, the slow processing of which is essential for efficient viral replication (Carrington et al., 1993; Schaad et al., 1996). The majority of full-length NIa is found in the nucleus of virus-infected cells (Carrington et al., 1993). However, NIa can also exist as a polyprotein with the 6K2 protein located upstream of NIa in the viral polyprotein. The 6K2 protein impedes nuclear localization of the 6K2-NIa polyprotein (Restrepo-Hartwig and Carrington, 1992) and directs NIa to cytoplasmic membrane vesicles, the sites of viral replication (Beauchemin et al., 2007; Cotton et al., 2009; Restrepo-Hartwig and Carrington, 1994; Schaad et al., 1997a). Many lines of evidence suggest that NIa is part of the viral replication complex involving several viral and host proteins (Beauchemin and Laliberte, 2007; Fellers et al., 1998; Li et al., 1997; Murphy et al., 1996; Puustinen

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and Ma¨kinen, 2004; Schaad et al., 1996). The VPg domain has NTP binding activity, is uridylylated by NIb, and may act as a primer for synthesis of viral RNA (Anindya et al., 2005; Murphy et al., 1996; Puustinen and Ma¨kinen, 2004; Schaad et al., 1996). In addition, VPg is involved in viral cell-to-cell and long-distance movement (Nicolas et al., 1997; Rajama¨ki and Valkonen, 1999, 2002; Schaad et al., 1997b). The NLSs and NoLS of potyviral NIa map to the N-proximal part of VPg (Carrington et al., 1991; Rajama¨ki and Valkonen, 2009; Schaad et al., 1996). The signal controlling nuclear localization of NIa is bipartite (Carrington et al., 1991; Rajama¨ki and Valkonen, 2009; Schaad et al., 1996). The regions of NIa controlling nucleolar targeting of NIa have been studied in PVA and found to map to the same regions as the NLSs. However, mutations in both NLS regions are required to abolish nuclear targeting of PVA NIa, whereas mutations in either NLS prevent nucleolar localization of NIa (Rajama¨ki and Valkonen, 2009). The most N-terminal NLS controls also localization of PVA NIa to CBs (Rajama¨ki and Valkonen, 2009). Mutations that affect nuclear localization of NIa are detrimental for genome amplification of TEV and PVA (Rajama¨ki and Valkonen, 2009; Schaad et al., 1996), suggesting that nuclear/nucleolar localization of NIa has an important role in virus infection. Potyviral NIa (or VPg) has been shown to interact with several host proteins (Dunoyer et al., 2004; Huang et al., 2010; Le´onard et al., 2004; Rajama¨ki and Valkonen, 2009; Schaad et al., 2000; Thivierge et al., 2008; Wittmann et al., 1997). The structure of VPg is intrinsically disordered (Grzela et al., 2008; Rantalainen et al., 2008), which may provide flexibility for many types of interactions. One of the interaction partners of VPg and/or NIa is the eukaryotic translation initiation factor eIF4E or its isoform (Robaglia and Caranta, 2006; Schaad et al., 2000; Wittmann et al., 1997) and the interaction appears important for virus infectivity (Le´onard et al., 2000; Robaglia and Caranta, 2006). Remarkably, interaction of TuMV NIa with eIF(iso)4E and the poly(A) binding protein 2 (PABP2) has been detected in the nucleus and nucleolus using bimolecular fluorescence complementation (BiFC) and colocalization experiments (Beauchemin and Laliberte, 2007; Beauchemin et al., 2007). By contrast, interaction with the 6K2-NIa polyprotein targets eIF(iso)4E and PABP2 to cytoplasmic membrane vesicles (Beauchemin and Laliberte, 2007; Beauchemin et al., 2007). The data suggest that these host proteins are needed in different viral processes. In healthy plants, most of PABP is cytoplasmic but some of the protein is redistributed to the nucleolus probably by the NIa (Beauchemin and Laliberte, 2007). NIa appears to mediate also eIF (iso)4E localization to the nucleolus (Beauchemin et al., 2007) and could interact simultaneously with eIF(iso)4E and PABP2 as shown by in vitro binding assays and BiFC and colocalization experiments (Beauchemin and Laliberte, 2007). Hence, all three proteins may be part of the same

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complex. Although the role of interaction between NIa and eIF(iso)4E and/or PABP2 in the nucleus is currently unclear, some possibilities can be suggested. The nuclear pool of eIF4E takes part in mRNA export from the nucleus but may also be involved in nuclear translation and NMD of RNA (Strudwick and Borden, 2002). PABP regulates the initiation of protein synthesis, nuclear export of mature mRNAs, and mRNA stability and decay (Mangus et al., 2003). Interaction with NIa might disrupt some of these functions. As mentioned above, some animal viruses are known to modulate and inhibit activities of translation initiation factors in order to favor viral replication and translation (reviewed by Thompson and Sarnow, 2000). Recent data connect nuclear/nucleolar targeting of plant viral proteins to interference with host antiviral defense. For example, the potyviral VPg protein has been observed to accumulate in the companion cells of upper leaves of a wild potato species ahead of virus infection, which suggested that VPg might interfere with host defense and hence facilitate infection of the cells that receive the virus via systemic transport (Rajama¨ki and Valkonen, 2003). Indeed, this hypothesis gained support in the experiments in which overexpression of VPg in leaf tissues temporarily interfered with cosuppression of gene silencing (i.e., RNA silencing), whereas NLS mutants of VPg, which exhibited reduced nuclear and nucleolar localization, were not able to suppress RNA silencing (Rajama¨ki and Valkonen, 2009). RNA silencing is a sophisticated, sequence-specific RNA degradation mechanism operating in the cytoplasm (Ruiz-Ferrer and Voinnet, 2009). A key feature of the RNA silencing pathway is the generation of dsRNA that corresponds in sequence to the target (virus) RNA. This dsRNA is cleaved into siRNAs by DCLs and these mediate the target specificity for RNA degradation (for reviews, see Carrington, 2000; Ruiz-Ferrer and Voinnet, 2009; Vance and Vaucheret, 2001; Voinnet, 2001). RNA silencing is a natural anti-viral defense system of plants but is also involved in gene regulation in a wide range of developmental and pathogen defense processes (Ruiz-Ferrer and Voinnet, 2009). To combat host defense RNA silencing, most plant viruses encode silencing suppressor proteins. The potyviral helper-component proteinase (HC-Pro) was the first detected viral suppressor of RNA silencing (Anandalakshmi et al., 1998; Brigneti et al., 1998; Kasschau and Carrington 1998) and the potyviral P1 protein may enhance its activity (Rajama¨ki et al., 2005). HC-Pro acts in the cytoplasm and, therefore, the association of VPg with RNA silencing suppression when localized in the nucleolus is intriguing because many host proteins involved in RNA silencing as well as the processing centers for small RNAs are located in the nucleus, nucleolus, and CBs as discussed in Section II (Pontes and Pikaard, 2008). In contrast, nuclear/nucleolar localization of the P19 protein of TBSV, tombusvirus, mediated by plant ALY proteins (mRNA-processing and

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-export factors) may be a defense mechanism of the plant to down-regulate the silencing suppressor activity of P19 (Canto et al., 2006). Previously, suppression of RNA silencing has been found to be connected to nuclear localization of another viral protein: the 2b protein, silencing suppressor of CMV. CMV 2b localizes in the nucleus and the nucleolus where it interacts with Argonaute 1 (AGO1), the core component of the RNA-induced silencing complex (Gonza´lez et al., 2010; Lucy et al., 2000; Zhang et al., 2006). More recently, it has also been shown that CMV 2b interacts with AGO4 (Gonza´lez et al., 2010). However, these interactions are not sufficient for suppression of RNA silencing, and hence their biological relevance remains so far unclear (Gonza´lez et al., 2010). The VPg of PVA interacts with fibrillarin in the nucleolus and CBs as detected by BiFC experiments (Rajama¨ki and Valkonen, 2009). Depletion of fibrillarin reduces PVA accumulation in Nicotiana benthamiana, suggesting a role for fibrillarin in virus infection (Rajama¨ki and Valkonen, 2009). As mentioned above, in GRV, fibrillarin is recruited for viral long-distance transport (Canetta et al., 2008; Kim et al., 2007a,b), but in potyviruses the role is likely to be different as long-distance transport of PVA is not compromised by depletion of fibrillarin (Rajama¨ki and Valkonen, 2009). Taking into account the silencing suppression activity of VPg, it could be suggested that VPg-fibrillarin might contribute to such an activity. Alternatively, the fibrillarin–VPg interaction may disrupt certain nucleolar functions (e.g., host transcription or pre-mRNA processing), which might explain the observed shutdown of host gene expression during potyvirus infection (Wang and Maule, 1995). In mammals, the IFN pathway plays a key role in the innate antiviral immune response whereas involvement of RNA silencing is still controversial (reviewed by Hale et al., 2008). However, emerging evidence suggests some parallels in how plant and animal viruses could use the nucleolus to counteract host defense. For example, the NS1 protein of influenza A virus is an important factor in counteracting IFN-based cellular antiviral mechanisms (Hale et al., 2008). In addition, NS1 is also able to suppress RNA silencing in plant, insect, and mammalian cells (Bucher et al., 2004; de Vries et al., 2009; Delgadillo et al., 2004; Haasnoot et al., 2007; Li et al., 2004). Remarkably, NS1 localizes to the nucleolus and interacts with nucleolin (Murayama et al., 2007).

D. Nucleolar localization of viral proteins for as yet unknown reasons NIb is a viral RNA-dependent RNA polymerase of potyviruses and hence involved in the replication complex (Hong and Hunt, 1996; Schaad et al., 1997a). While viral replication takes place in the cytoplasm on cellular

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membranes, NIb is also targeted to the nucleus and the nucleolus (Baunoch et al., 1991; Restrepo et al., 1990). Nuclear targeting of NIb appears to be highly sensitive to alterations in protein conformation and is eliminated by deletions, dipeptide insertions and amino acid substitutions introduced also into parts of NIb other than the NLSs (Li and Carrington, 1993). Nuclear and nucleolar localization of NIb may be important for the viral infection cycle, because mutations in the NLSs abolish infectivity of TEV (Li and Carrington, 1995; Li et al., 1997). Three host proteins, the PABP2, the heat shock cognate 70 protein (Hsc70-3), and the eukaryotic translation elongation factor 1A, eIF1A, interact with NIb (Dufresne et al., 2008; Thivierge et al., 2008; Wang et al., 2000b). However, none of these protein interactions was found in the nucleolus, shedding no light on the role of the nucleolar localization of NIb. Besides nuclear inclusion proteins, the P3 protein of TEV is targeted to the nucleus and nucleolus of virus-infected tobacco cells, as detected by immunogold labeling (Langenberg and Zhang, 1997). P3 is a nonstructural protein with no well-characterized function. However, it is involved in virus multiplication (Kekarainen et al., 2002) and virus–host interactions (Chu et al., 1997; Eggenberger et al., 2008; Jenner et al., 2003; Johansen et al., 2001). A role for nuclear/nucleolar localization of P3 is currently unknown. The multifunctional nucleocytoplasmic shuttling P6 protein encoded by plant para-retrovirus CaMV has also been found to localize to the nucleolus (Haas et al., 2005). Although a NoLS has not been identified for this protein, its nucleolar import might be facilitated by its interaction with ribosomal proteins of the 60S ribosomal subunit (Haas et al., 2008). The presence of P6 in the nucleolus, where assembly of ribosomal subunits occurs, raises the possibility that P6 might interact directly with ribosomes before their export to render them competent for translation of the CaMV polycistronic mRNA. Indeed, P6 interacts with the ribosomal proteins L18 (Leh et al., 2000), L24 (Park et al., 2001), and L13 (Bureau et al., 2004), which hence may be the potential targets because they participate in the formation of the 60S subunit in the nucleolus (Andersen et al., 2002). Another functional activity of CaMV P6 is suppression of RNA silencing. This protein interferes with RNA-directed RNA polymerase 6 (RDR6)dependent RNA silencing via inhibition of the dsRNA-binding protein DRB4, a protein normally enhancing DCL4-mediated dicing. However, unlike nuclear targeting, the nucleolar localization of P6 is completely dispensable for its silencing suppression function (Haas et al., 2008). Transport of the viral genome into the nucleus is an obligatory step in the replication cycle of plant DNA viruses such as the begomoviruses. CPs of monopartite begomoviruses (such as TYLCV and ToLCJAV) are nucleocytoplasmic shuttling proteins thought to be involved in this process (Rojas et al., 2001; Sharma and Ikegami, 2009). Interestingly, these

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proteins also contain NoLSs targeting them to the nucleolus (Sharma and Ikegami, 2009). However, the biological significance of nucleolar localization of begomoviral CP is unclear. NLSs have been identified in three of the seven proteins encoded by plant nucleorhabdovirus, MFSV. Remarkably, two of them, the nucleocapsid (N) protein and phosphoprotein (P), localize to the nucleolus but only when they are coexpressed in plant tissues (as fusions with GFP) using the Agrobacterium system; when expressed individually these proteins do not target nucleoli (Tsai et al., 2005). This clearly indicates the interdependent character of nucleolar targeting for the N and P proteins. However, the molecular mechanisms responsible for this effect and its biological relevance remain to be explored. Another plant virus protein with nucleolar localization is the CP of SPMV (satellite panicum mosaic virus) (Qi et al., 2008). Some of the functional and biochemical properties of the SPMV CP including the nucleolar association of SPMV CP, its RNA binding activity (Desvoyes and Scholthof, 2000), and the involvement of the CP in systemic movement (Omarov et al., 2005) are similar to those of GRV ORF3. By analogy therefore, the authors have suggested that nucleolar localization of SPMV CP may be essential for its role in the systemic movement of SPMV as in the case of GRV ORF3. Collectively, all of these findings support an active role of the nucleolus and fibrillarin in various aspects of the virus infection cycle and interactions with host cells promoting systemic infections with plant viruses belonging to various taxonomic groups.

E. Formation of viral ribonucleoprotein complexes (RNPs) in the nucleolus The nucleolus contains a complex machinery for rRNA modification and rRNP assembly and may provide an environment which allows other forms of functional RNPs, such as SRP, telomerase, splicing snRNPs, and viral RNPs to exploit the nucleolus or nucleolar components in their biogenesis pathways (reviewed by Boisvert et al., 2007). For example, fibrillarin is one of the four core protein components of a box C/D snoRNP complex (reviewed by Tran et al., 2004). Being a methyltransferase, fibrillarin (as a component of box C/D snoRNPs) functions in the 20 -in vitro-methylation and processing of pre-rRNA. In addition, human fibrillarin forms a subcomplex with splicing factor 2-associated p32, protein arginine methyltransferases, and tubulins a3 and b1 that is independent of its association with snoRNPs, suggesting that fibrillarin may also coordinate protein methylation during the processes of ribosome biogenesis (Yanagida et al., 2004). Furthermore fibrillarin interacts with some other cellular proteins such as SMN protein ( Jones et al., 2001) and the nuclear DEAD box protein p68, an RNA-dependent ATPase and RNA

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helicase (Nicol et al., 2000). However, the physiological role of these interactions is unclear and may be based on novel natural functions of fibrillarin that remain to be established. The most studied viral RNP complex containing fibrillarin is the GRV ORF3 complex with elements of regular helical structure that is capable of long-distance movement via the phloem. Assembly of GRV RNP particles occurs in the cytoplasm and requires fibrillarin relocalized from the nucleolus (Kim et al., 2007a,b; Taliansky et al., 2003; also see above). RNP particles similar in architecture and infectivity to the viral RNPs formed in vivo, have been reconstituted in vitro from fibrillarin, the ORF3 protein and viral RNA (Kim et al., 2007a,b). Taking the study further, the in vitro experiments have shown that the ORF3-fibrillarin interaction occurs between the leucine-rich region (L149 in particular) of the ORF3 protein and the GAR domain of fibrillarin (Kim et al., 2007b) known to be responsible for interaction with other proteins such as SMN ( Jones et al., 2001). This interaction leads to formation of single-layered ring-like complexes of ORF3 with fibrillarin as was shown by atomic force microscopy (Canetta et al., 2008). The diameter of these ORF3 protein-fibrillarin rings is 18–22 nm which correlates with that of the filamentous RNP particles (Canetta et al., 2008). It thus appears that the ORF3 protein fibrillarin rings interact with viral RNA, encapsidating it and reorganizing it into helical structures, and thereby play a key role in the assembly of umbraviral RNP complexes. These results demonstrate that, in addition to traditional functions in rRNA processing and modification, fibrillarin possesses completely novel functions in mediating assembly of umbraviral RNPs. These functions are presumably based on the ability of the ORF3 protein to interact and form ring-like complexes with fibrillarin such that the virus alters and exploits the properties of fibrillarin for successful virus propagation. Other viral proteins, such as the nucleocapsid protein encoded by porcine arterivirus also interact with fibrillarin in an RNA-dependent manner (Yoo et al., 2003). However, the structure and architecture of these complexes and how they impact the viral infection cycle remain unknown.

V. CONCLUSIONS AND PERSPECTIVES The nucleolus is a highly conserved feature of eukaryotic cells that has a key role as the site of ribosome subunit production. However, recent multiple lines of investigation have confirmed and characterized additional roles for nucleoli in other important cellular processes including cell-cycle control, stress responses, and coordination of the biogenesis of a number of other functional RNPs. The nucleus has also been shown to

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play a crucial role in the infection cycle of various viruses, and the nucleolar localization of viral proteins has recently been described as a pan-virus phenomenon (Hiscox, 2002, 2007). In this regard, plant viruses are not different from other eukaryotic viruses. The past few years have brought remarkable progress in our understanding of why and how some plant viruses (in particular, umbraviruses and potyviruses) target the nucleolus and the functional role of the interaction between viral and nucleolar proteins in the plant virus infection cycle. For many other plant virus proteins, nucleolar localization and interaction with nucleolar components have also been demonstrated, and functional implications of these findings is a challenge for future research. We anticipate that more information will emerge about the mechanisms involved in regulating nucleolar function and structure in response to plant virus infections and hijacking nucleolar functions for the virus infection cycle. There are now several examples in which the plant viruses also target other subnuclear bodies, such as CBs. In particular, for umbraviruses the role of CBs in nucleolar trafficking of the ORF3 protein has been established. The potential role of sub-nuclear structures in other plant virus infections will be addressed in the future. The study of viral interactions with the nucleolus also provides unique and valuable tools to gain new insights into novel nucleolar functions and processes. For example, as previously discussed, the major nucleolar protein fibrillarin, is involved in formation and long-distance movement of umbraviral RNP particles. These functions as well as other potential yet unrecognized natural functions of nucleolar proteins will be the focus of much future research. On a practical level, both the plant cell and viral biology of the nucleolus can, and hopefully will be exploited for the design of novel strategies to control plant virus infections.

ACKNOWLEDGMENTS This work was supported by Scottish Government Rural and Environment Research and Analysis Directorate (M.T. and J.W.S.B.), Academy of Finland (grants 118766 and 134759 to J.V), Ministry of Education and Science of Russian Federation (grant 02.740.11.5145 to M.T and N.O.K.) and Russian Foundation for Basic Research (grant RFBR-10-04-00522 to N.O. K).

REFERENCES Ahmad, Y., Boisvert, F.-M., Gregor, P., Cobley, A., and Lamond, A. I. (2008). NOPdb: Nucleolar proteome database – 2008 update. Nucleic Acids Res. 37:D181–D184. Amineva, S. P., Aminev, A. G., Palmenberg, A. C., and Gern, J. E. (2004). Rhinovirus 3C protease precursors 3CD and 3CD’ localize to the nuclei of infected cells. J. Gen. Virol. 85:2969–2979.

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INDEX A Alphapapillomavirus oncogenicity, HPV biochemical functions, 55 OT and NOT types, 53–54 p53 and PDZ degradation activities, 54–55 Antibody-dependent enhancement (ADE), 7, 19 Antigenic drift and shift, 66–69 B Bird virus, influenza highly pathogenic avian (HPAI) H5N1 virus, 72–73 H9N2 viruses, 73–74 vaccines production, 73 Blood and peripheral tissues, poliovirus host barriers, 97–98 transient viremia, 96–97 C Cajal bodies (CBs), 126 Cell cycle, DENV replication ADE, 19 cell receptors, 18 characteristics and functions, 12–16 endosome membrane, adsorption and fusion, 19 E protein, 11–18 genome expression, 20–21 replication, 21–24 GRP78, adaptor proteins, 18–19 intracellular transport, 19–20 maturation and release encapsulation, 24 particles maturation, 25 prM cleavage, 25 V-ATPases, 25–26 viral RNA-C protein assembly, 24–25 schematic representation, 17 Central nervous system (CNS), poliovirus

host barriers non-neuroinvasive variant, 101 provocation poliomyelitis, 102–103 viral trafficking, neural route, 101–102 invasion blood–brain barrier, 98–99 neurons, 99–100 Cervical carcinogenesis, 44–45 Coxsackievirus B3 (CVB3), 88 D Dengue virus (DENV) replication cell cycle ADE, 19 cell receptors, 18 characteristics and functions, 12–16 endosome membrane, adsorption and fusion, 19 E protein, 11–18 genome expression and replication, 20–24 GRP78, adaptor proteins, 18–19 intracellular transport, 19–20 maturation and release, 24–26 schematic representation, 17 disease agent, 4–5 clinical manifestations, 5–6 eco-epidemiology, 5 pathogenesis, 6–8 genome and proteins 5’ and 3’ untranslated regions (UTRs), 10–11 circularization, 9 NS5 and NS3 proteins, 10 translational cleavage, 11 viral polyprotein, 8–10 history, 3–4 siRNAs and replication control, 26–27 Dense fibrillar component (DFC), 123 DENV replication. See Dengue virus (DENV) replication

159

160

Index

E E6 and E7 viral oncogenes, HPV cell polarity, 49 hypophosphorylated pRb, 49–51 immune response, 51 interaction partners, 50 mutation rate, 53 p53 degradation, 49 protein structures, 51–53 Enterovirus 71 (EV71), 88 E protein, 11–19 F Fibrillar centers (FC), 123 G Gastrointestinal tract route, poliovirus CD155 receptor, 88–89 host barriers bile salts and digestive enzymes, 92 cell surface, 95 defense mechanisms, 94 gut architecture and natural homeostasis, 96 immunoglobulin A and CD155 receptor, 94 innate immune response, 95–96 mucus layer lines, 92 stomach acid, 91–92 viral population diversity assay, 93 intestinal mucosa, 89, 90 oropharynx, viral replication, 89 shedding and transmission, 91 viral infection, 89–91 virus transmission, 88 Genome, DENV replication expression functions, 20 viral protein synthesize, 21 replication autophagy, 23–24 mutation, 22–23 RNA synthesis, 21–22 viral RNA circularization, 23 Granular component (GC), 123 GRP78, adaptor proteins, 18–19 H Hemagglutinin (HA) antigen, 65–66, 73 Highly pathogenic avian (HPAI) H5N1 virus, 72–73

HIV, 130–131 H9N2 viruses, 73–74 Human papillomavirus (HPV) alphapapillomavirus oncogenicity biochemical functions, 55 OT and NOT types, 53–54 p53 and PDZ degradation activities, 54–55 cervical carcinogenesis and, 44–45 classification, 42 clinical implications b-and g-PVs genus, 43 Bayesian phylogenetic tree, 42, 43 a-PV genus, 43–44 E6 and E7 viral oncogenes cell polarity, 49 hypophosphorylated pRb, 49–51 immune response, 51 interaction partners, 50 mutation rate, 53 p53 degradation, 49 protein structures, 51–53 life cycle epithelial infection, 45–46 productive infection, 46–47 structure and organization, 45 viral carcinogenicity cervicovaginal epithelium infection, 47 phylogenetic approaches, 48–49 I Inactivated vaccines (INV), influenza, 69 Influenza a2–6 and 3 linkages, 66 antigenic drift and shift, 66–69 bird viruses highly pathogenic avian (HPAI) H5N1 virus, 72–73 H9N2 viruses, 73–74 vaccines production, 73 generating process, 70 hemagglutinin (HA) and neuraminidase (NA) protein, 65–66 inactivated vaccines (INV), 69 live-attenuated vaccines (LAV), 69–70 Orthomyxoviridae family, 65 pandemic vaccine H1N1 vaccine production, 71–72 outcomes, 72 respiratory illness cases, 71 proteins, 66 respiratory disease, 64

Index

seasonal epidemics, 64 universal vaccine, 76–77 vaccine efficacy, 70 vaccines future cell-based cultures, 74–76 drawbacks, 74–75 efficacy improvement, 76 virion structure, 65 In situ hybridization, 135 Internal ribosome entry sites (IRESs), 130 L Lactoferrin, 94 Lipid droplets (LDs), 24 Live-attenuated vaccines (LAV), influenza, 69–70 M Muller’s ratchet, 106 N Neuraminidase (NA) protein, 65–66 Nonsensemediated decay (NMD), 127 Nuclear export protein (NEP), 66 Nucleolus plant virus proteins host antiviral defense, 140–143 nucleolar localization, 143–145 nucleolar trafficking, 132 viral ribonucleoprotein complexes (RNPs) formation, 145–146 viroids and viruses replication, 134–135 virus movement, 135–139 ribosomal RNA (rRNA), 122 structure and functions B23, 125 Cajal bodies (CBs), 126 fibrillarin, 124–125 nuclear pore complexes, 125 nucleolin, 125 nucleus, 123 plant nucleolus, 127–128 protein composition characterization, 126–127 regions, 123–124 ribosomal subunits productions, 126 transcription and precursor rRNAs processing, 124 viruses interactions cytoplasm, 129–131 nucleus, 129 virus-host interactions, 121–122

161

O Open reading frame (ORF), 9–10 P Plant nucleolus, structure and functions biochemical fractionation, 127–128 functions, 127 mRNAs, 128 RNA and RNP maturation pathway, 128 Plant virus proteins fibrillarin interaction, 132–134 host antiviral defense, nucleolar targeting fibrillarin, 143 IFN pathway, 143 nuclear inclusion protein a and b (NIa/NIb), 140–141 potyviral replication-associated proteins, 140 RNA silencing, 142 VPg protein, 141–142 nucleolar localization, 143–145 viral ribonucleoprotein complexes (RNPs) formation, 145–146 viroids and viruses replication, nucleolus, 134–135 virus movement poleroviruses, 139 systemic spreading stages, 136 triple gene block protein (TGB), 139 umbraviruses, 136–139 Poleroviruses, 139 Poliovirus effects, 86 research benefit, 87–88 route and barriers blood and peripheral tissues, 96–98 CNS, 98–103 gastrointestinal tract, 88–96 vaccines, 86–87 viral population, host barriers effects excessive mutagenesis, 105 fitness loss, limited diversity, 105–107 RNA virus mutations, 104–105 Poly(A)-binding protein (PABP), 20 R Ribonucleoprotein complex (RNP), 66 RNA-dependent RNA polymerases (RdRps), 104 RNA virus mutations, 104–105 Route and barriers, poliovirus

162

Index

Route and barriers, poliovirus (cont.) blood and peripheral tissues host barriers, 97–98 transient viremia, 96–97 CNS host barriers, 101–103 invasion, 98–100 gastrointestinal tract CD155 receptor, 88–89 host barriers, 91–96 intestinal mucosa, 89, 90 oropharynx, viral replication, 89 shedding and transmission, 91 viral infection, 89–91 virus transmission, 88 S Satellite panicum mosaic virus (SPMV), 145

U Umbraviruses Cajal body-like structures (CBL) formation, 137 fibrillarin, 138–139 GRV infection biology, 137 nucleolar import pathway, 138 ORF3 protein, 137 proteins, 136–137 Untranslated regions (UTRs), 10–11 V Viral carcinogenicity, HPV cervicovaginal epithelium infection, 47 phylogenetic approaches, 48–49 Viroid RNAbinding protein 1 (VIRP1), 135