Porcine Viruses : From Pathogenesis to Strategies for Control [1 ed.] 9781910190920, 9781910190913

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Porcine Viruses From Pathogenesis to Strategies for Control

Edited by

Hovakim Zakaryan Caister Academic Press

Porcine Viruses

From Pathogenesis to Strategies for Control

https://doi.org/10.21775/9781910190913

Edited by Hovakim Zakaryan Group of Antiviral Defense Mechanisms Institute of Molecular Biology of the National Academy of Sciences Yerevan Armenia

Caister Academic Press

Copyright © 2019 Caister Academic Press Norfolk, UK www.caister.com British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-1-910190-91-3 (paperback) ISBN: 978-1-910190-92-0 (ebook) Description or mention of instrumentation, software, or other products in this book does not imply endorsement by the author or publisher. The author and publisher do not assume responsibility for the validity of any products or procedures mentioned or described in this book or for the consequences of their use. 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 permission of the publisher. No claim to original U.S. Government works. Ebooks Ebooks supplied to individuals are single-user only and must not be reproduced, copied, stored in a retrieval system, or distributed by any means, electronic, mechanical, photocopying, email, internet or otherwise. Ebooks supplied to academic libraries, corporations, government organizations, public libraries, and school libraries are subject to the terms and conditions specified by the supplier.

Contents Prefacev 1

African Swine Fever Virus

2

Classical Swine Fever Virus

21

3

Foot-and-Mouth Disease Virus

43

4

Porcine Circoviruses

81

5

Porcine Epidemic Diarrhoea Virus

107

6

Porcine Parvovirus

135

7

Porcine Reproductive and Respiratory Syndrome Virus

149

8

Swine Vesicular Disease Virus

181

Erik Arabyan, Armen Kotsinyan, Astghik Hakobyan and Hovakim Zakaryan Sandra Blome

Francisco Sobrino, Flavia Caridi, Rodrigo Cañas-Arranz and Miguel Rodríguez-Pulido Sheela Ramamoorthy and Pablo Piñeyro Changhee Lee

André Felipe Streck and Uwe Truyen Alexander N. Zakhartchouk, Sujit K. Pujhari and John C.S. Harding

Estela Escribano-Romero, Miguel A. Martín-Acebes, Angela Vázquez-Calvo, Emiliana Brocchi, Giulia Pezzoni, Francisco Sobrino and Belén Borrego

1

Index199

Current Books of Interest The Prion Protein2019 Plant Genomics2019 Methylotrophs and Methylotroph Communities2019 Microbial Ecology: Current Advances from Genomics, Metagenomics and Other Omics2019 Plant-Microbe Interactions in the Rhizosphere2018 Prions: Current Progress in Advanced Research (Second Edition)2019 Microbiota: Current Research and Emerging Trends2019 Lactobacillus Genomics and Metabolic Engineering2019 Cyanobacteria: Signaling and Regulation Systems2018 Viruses of Microorganisms2018 Protozoan Parasitism: From Omics to Prevention and Control2018 Genes, Genetics and Transgenics for Virus Resistance in Plants2018 DNA Tumour Viruses: Virology, Pathogenesis and Vaccines2018 Pathogenic Escherichia coli: Evolution, Omics, Detection and Control2018 Postgraduate Handbook: A Comprehensive Guide for PhD and Master’s Students and their Supervisors2018 Enteroviruses: Omics, Molecular Biology, and Control2018 Molecular Biology of Kinetoplastid Parasites2018 Bacterial Evasion of the Host Immune System2017 Illustrated Dictionary of Parasitology in the Post-Genomic Era2017 Next-generation Sequencing and Bioinformatics for Plant Science2017 The CRISPR/Cas System: Emerging Technology and Application2017 Brewing Microbiology: Current Research, Omics and Microbial Ecology2017 Metagenomics: Current Advances and Emerging Concepts2017 Bacillus: Cellular and Molecular Biology (Third edition)2017 Cyanobacteria: Omics and Manipulation2017 Brain-eating Amoebae: Biology and Pathogenesis of Naegleria fowleri2016 Foot-and-Mouth Disease Virus: Current Research and Emerging Trends2017 Staphylococcus: Genetics and Physiology2016 Chloroplasts: Current Research and Future Trends2016 Microbial Biodegradation: From Omics to Function and Application2016 www.caister.com

Preface

For humans, pork meat is one of the most complete dietary sources of protein. According to the Food and Agriculture Organization of the United Nations, animal protein production will grow three times by 2050, and meat production, including of pork, will double. An increase in intensification is necessary, because arable land cannot be increased in proportion. However, viral diseases cause significant animal losses and represent a major threat to global pig farming industry. Therefore, the understanding of molecular biology, pathogenesis, host–virus interaction and epidemiology of these viruses is essential for reducing the burden of viral outbreaks. The overall aim of this book is to review the most important and dangerous porcine viruses that have emerged in the global swine population. It covers different DNA and RNA viruses about which we have learnt much during the last decades. In Chapter 1, my colleagues and I discuss African swine fever virus (ASFV), the causative agent of highly lethal haemorrhagic fever of domestic pigs and wild boar. It is a large, enveloped, double-stranded DNA virus that is the only known DNA arbovirus since it is transmitted by soft ticks of the genus Ornithodoros. Most recently, ASFV was introduced into Georgia and then spread to the Russian Federation, Belarus, Ukraine, and some EU member states. In the absence of effective vaccines and antiviral drugs, this virus continues to pose a global risk for pig industry. Chapter 2 is about classical swine fever virus (CSFV), a small enveloped RNA virus of the genus Pestivirus in the Flaviviridae virus family. Although live attenuated vaccines are currently available, it remains a major threat to profitable pig production worldwide because CSFV infection is still associated with high mortality rates. Sandra Blome summarizes CSFV properties, pathogenesis, clinical picture and control options. Chapter 3 focuses on foot-and-mouth disease virus (FMDV), which is the prototypic member of the Picornaviridae family. It causes an acute systemic vesicular disease affecting livestock worldwide. Francisco Sobrino and colleagues discuss different aspects of FMDV infection, including new strategies for viral control by vaccination and other antiviral strategies. This chapter also covers the current approaches for virus diagnosis. In Chapter 4, Sheela Ramamoorthy and Pablo Piñeyro provide an overview of porcine circoviruses, focusing particularly on porcine circovirus strain 2 (PCV2), which was first isolated in 1997. PCV2 is a single-stranded DNA virus belonging to the Circoviridae family. PCV2 infection by itself causes only mild disease but co-factors such as other infections are involved in the development of severe diseases. Molecular biology, pathogenesis, immune response, diagnosis and control strategies are discussed in detail. Chapter 5 is about porcine epidemic diarrhoea virus (PEDV), the aetiological agent of severe diarrhoea and dehydration. Although this virus was first reported in

vi

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Preface

Europe, it has become problematic in Asian countries such as China, Japan, Thailand and the Philippines. Owing to high morbidity and mortality in piglets, PEDV has a substantial economic burden in affected countries. Changhee Lee discusses molecular and cellular biology of the virus, as well as diagnostic procedures, epidemiology and control strategies. In Chapter 6, André Felipe Streck and Uwe Truyen describe the biology, pathogenic potential and strain variation of porcine parvovirus (PPV). This virus is considered the main cause of reproductive disorders in pigs with no maternal clinical signs. PPV is a small, non-enveloped, single-stranded DNA virus belonging to the Parvoviridae family. Although losses are low in vaccinated herds, PPV can cause devastating abortion storms in unvaccinated herds, or in those herds, where new antigenic types are circulating. Chapter 7 is devoted to porcine reproductive and respiratory syndrome virus (PRRSV), a small enveloped RNA virus belonging to the Arteriviridae family. PRRSV causes reproductive failure in herds and respiratory tract illness in young pigs. In this chapter, Alexander Zakhartchouk and colleagues summarize the current understanding of PRRSV, including the virus molecular biology, virus–host cell interactions, pathogenesis, diagnostic procedures and epidemiology. They also provide an overview of currently available vaccines and a novel vaccine development. The last chapter (Chapter 8) is about swine vesicular disease virus (SVDV), which belongs to the Enterovirus genus within the Picornaviridae family. SVDV is genetically highly related to the human coxsackie virus B5. It causes a vesicular disease with clinical signs similar to those of foot-and-mouth disease. Francisco Sobrino, Belén Borrego and colleagues discuss different aspects essential for understanding the infectious cycle of SVDV. They also provide an overview of current strategies for SVDV control by vaccination and other antiviral strategies. It took about a year to complete this book, during which time the chapters were written, edited and re-edited in order to improve and present high-quality reading material. My primary acknowledgement must go to all our contributing authors for their significant work, enthusiasm and cooperation. In addition, I thank Annette Griffin from Caister Academic Press for her great assistance and patience during all this time. Finally, but most importantly, I wish to extend appreciation to my family members, particularly to my beautiful wife, Kamila, for her invaluable support throughout the writing and editing process. Dr Hovakim Zakaryan

African Swine Fever Virus Erik Arabyan, Armen Kotsinyan, Astghik Hakobyan and Hovakim Zakaryan*

1

Group of Antiviral Defense Mechanisms, Institute of Molecular Biology of the National Academy of Sciences, Yerevan, Armenia. *Correspondence: [email protected] https://doi.org/10.21775/9781910190913.01

Abstract African swine fever virus (ASFV) is a large DNA virus belonging to the family Asfarviridae. It is the causative agent of an acute haemorrhagic fever in domestic pigs and wild boar with high fatality rates, up to 100%, in the acute forms. The disease is currently present in Africa and Europe, where it causes a high socio-economic impact. In this chapter we have addressed different aspects of ASFV including the biology of the virus which is relevant for understanding the viral disease. We have also discussed the pathological changes caused by high and low virulence ASFV strains. Finally, this chapter addresses the current approaches for ASFV diagnosis, as well as presents an overview of research efforts towards the development of effective vaccines during the past decades. As an alternative to vaccine development, the current state in antiviral research is also presented. History The disease caused by African swine fever virus (ASFV) was first observed in settlers’ pigs in Kenya in 1909 and was reported by Montgomery (1921) as an acute haemorrhagic disease distinct from classical swine fever. After the first report, the disease was also observed in South Africa and Angola (Gago da Câmara, 1932; Steyn, 1932). Although Gago da Câmara determined that the disease observed in pigs was not erysipelas, the viral identity was confirmed in 1943 (Mendes, 1994). Interestingly, pigs belonging to the indigenous population of Angola demonstrated increased resistance to African swine fever (ASF) suggesting that they served as a source of infection for the pigs farmed by European settlers (Mendes, 1994). During the same period, the link between warthogs and ASF was shown (Montgomery, 1921). However, the possible involvement of an argasid tick vector in the epidemiology of ASF was confirmed much later, in 1960s, when Sánchez Botija (1963) reported about the argasid tick Ornithodoros erraticus in the epidemiology of ASF in Spain. Studies undertaken in southern and eastern Africa revealed ASFV in O. moubata complex ticks inhabiting both pig shelters and warthog burrows (Penrith et al., 2004).

2  | Arabyan et al.

The first cases of ASF outside of Africa was reported in Portugal in 1957 and again in 1960 with its further establishment in the Iberian Peninsula for many decades (Wilkinson, 1984). ASF has also caused sporadic outbreaks in other European countries, such as France (1964, 1967 and 1974), Italy (1967, 1969 and 1993), Andorra (1975), Belgium (1985), Malta (1978), and The Netherlands (1986). However, in 1995, the disease was eradicated from the continental Europe, except Sardinia, where it has been endemic since 1978. It is the only territory beyond Africa that is considered endemic for ASF at the present time. In 2007, ASF re-emerged in Europe for a second time, following a single introduction into Georgia in the Caucasus, from which it spread rapidly to the Russian Federation, Armenia, Azerbaijan and a number of countries in Eastern Europe, including some EU countries (Estonia, Lithuania, Latvia, Poland, the Czech Republic and Romania). Owing to the lack of vaccines or antiviral drugs, as well as in a context of increased trade activity, there is widespread concern that ASF may spread beyond affected countries. Classification ASFV was initially classified as an iridovirus, based mainly on the similarity of virion morphologies. However, increasing knowledge of ASFV biology, particularly genome analysis, led to its reclassification as a new DNA virus family, Asfarviridae (Asfar, African swine fever and related viruses) (Penrith et al., 2004). Until recently, ASFV was considered to be the only known virus of this family. In 2009, the complete sequence of the type B DNA polymerase (PolB) gene of Heterocapsa circularisquama DNA virus revealed a close link with Asfarviridae, suggesting a co-occurrence of marine and terrestrial viruses in the same family (Ogata et al., 2009). In 2015, La Scola and colleagues isolated eight strains of a new giant virus named faustovirus from Vermamoeba vermiformis (Reteno et al., 2015). The genomes of faustoviruses showed that they are related to ASFV, although the faustovirus gene repertoire is three times larger than that of ASFV. Two other viruses, kaumoebavirus and pacmanvirus, were branched within the cluster encompassing ASFV and faustovirus (Bajrai et al., 2016; Andreani et al., 2017) (Fig. 1.1). Thus, these findings and future viruses may deeply alter the current knowledge of Asfarviridae family in many ways. The comparison of ASFVs based on the partial sequence of B646L gene has allowed the differentiation of ASFV isolates collected worldwide. Using results from partial sequencing, phylogenetic analysis has divided ASFV isolated into 23 genotypes (Bastos et al., 2003). The newest genotype was reported in Ethiopia (Achenbach et al., 2017). ASFV is also classified as the only known arbovirus (arthropod-borne virus) with a DNA genome, since soft ticks are involved in the sylvatic transmission cycle of ASFV in Africa (O. moubata) and in Europe (O. erraticus). Molecular biology of the virus Structure of ASFV particles The ASFV particle has an icosahedral shape with an average size of 200 nm. Two-dimensional analysis of ASFV purified by Percoll gradient centrifugation has identified 54 structural proteins with different molecular weights (Esteves et al., 1986). These proteins are localized in several concentric domains: an internal core containing viral DNA and nucleoid coated by a

African Swine Fever Virus |  3 Methanohalophilus mahii, Euryarchaeota Faustovirus 100/1/100 Faustovirus ST1

99.9/1/100

Pacmanvirus A23 47.5/0.655/57 Heterocapsa ciruclarisquama DNA virus 01

98/1/86 Kaumoebavirus

99.9/1/100

99.8/1/100 Benin 97/1 85.6/0.955/74

ASFV

Georgia 2007/1

Ken06.Bus Klebsiella pneumoniae UHKPC27, Gammaproteobacteria 0.4

Figure 1.1  Phylogenetic tree based on the amino acid sequence alignment of DNA polymerase beta of viruses from Asfarviridae family.

thick protein layer (core shell); an inner lipid envelope; and the capsid (Andrés et al., 1997). The extracellular ASFV particles possess an external envelope acquired by budding from the plasma membrane. Although the external envelope is similar to the plasma membrane, viral proteins, p12, p24 and pE402R, have been reported to localize at the outer envelope (Sanz et al., 1985; Carrascosa et al., 1993; Rodríguez et al., 1993). The capsid is composed of about 2000 capsomers arranged in a hexagonal lattice. Computer-filtered electron micrographs have revealed capsomers with a hexagonal outline and a hole in the centre. The intercapsomer distance varies from 7.4 to 8.1 nm (Carrascosa et al., 1984). Protein p72, encoded by viral B646L gene, is the main component of capsomers, accounting for about one third of the protein mass of ASFV (Carrascosa et al., 1986). It has been shown that the correct folding of p72 requires the expression of another viral protein, pB602L, which acts as a molecular chaperone (Epifano et al., 2006a). Another component of the capsid is pE120R whose incorporation into the virus particle is dependent on p72 expression. This protein is involved in the microtubule-mediated transport of ASFV particles from the viral factories to the plasma membrane (Andrés et al., 2001). The lack of structural protein pB438L leads to the formation of viral particles with a tubular structure, suggesting that it is likely to be involved in the stabilization of the icosahedral vertices of ASFV particles (Epifano et al., 2006b). In addition, the virion contains proteins for the transcriptional machinery, capping, polyadenylation, nucleoside triphosphate phosphohydrolases and nucleoproteins such as the DNA-binding protein p10 and pA104R, which is similar to the histone-like proteins of bacteria (Salas, 1999; Andrés et al., 2002).

4  | Arabyan et al.

Viral genome structure The viral genome is a linear double-stranded DNA molecule that varies in length between different isolates from 170 to 193 kbp (Chapman et al., 2008). The differences in genome length are due to gain or loss of open reading frames (ORF) from the multigene families (MGF) encoded by virus. ASFV contains five MGFs (MGF 100, 110, 300, 360 and 505/530,) named to reflect the average number of codons present in each gene. Comparison of the genome sequences of ASFV isolates revealed multiple gene deletions in MGF 110 and 360 (Chapman et al., 2008). Moreover, repeated passages of field isolates through cell cultures result in the loss of genes of the MGF 110 family, suggesting that these genes may determine the virulence of ASFV in different hosts (Pires et al., 1997). The ASFV genome contains a conserved, centrally located genomic core (125 kbp) in which major changes are rare and high variability sequences confined to the left (38 to 47 kbp) and right (13 to 16 kbp) terminal regions (Fig. 1.2). Both strands of DNA encode genes that are closely spaced and do not contain introns. Promoter sequences are short and A + T rich. The genome termini are covalently cross-linked hairpin loops presented in two forms, inverted and complementary to each other (González et al., 1986). Cellular biology of the virus Viral entry The viral cycle starts with the infection of monocytes and macrophages, the natural target cells of ASFV in pig. The restricted cell tropism of ASFV indicates that a monocyte/ macrophage-specific receptor is required for infection. CD163 is the high affinity scavenger receptor whose expression is specific for the cells of monocyte/macrophage lineage. It has been shown to be important for ASFV entry, since incubation of macrophages with a specific anti-CD163 antibody decreases the level of infection in a dose-dependent manner (Sánchez-Torres et al., 2003). However, more recent studies have reported that pigs possessing a complete knockout of CD163 by CRISPR/Cas9 are not resistant to infection with ASFV Georgia2007/1 strain (Popescu et al., 2017), thereby suggesting that other macrophage surface proteins are also involved in the infection process. On the other hand, viral proteins responsible for ASFV binding to host cells have been described. Neutralizing antibodies against viral proteins p12, p32, p54 and p72 inhibited both ASFV attachment to and internalization into Vero cells and macrophages, indicating that these proteins are involved in the early stages of ASFV infection (Carrascosa et al., 1991; Gomez-Puertas et ASFV BA71V Genome Left variable region

Genomic core

Genes: MGF300; MGF360; MGF505/530

Right variable region

Genes: MGF100; MGF360; MGF505/530

Figure 1.2  Genomic regions of ASFV BA71V strain. Left and right variable regions contain multigene families (MGFs).

African Swine Fever Virus |  5

al., 1996). Interestingly, baculovirus-expressed p32, p54 and p72 proteins were not able to protect pigs against the virulent virus (Neilan et al., 2004). This result suggests that other viral proteins should be involved in ASFV entry, and therefore further studies may define both viral and cellular proteins responsible for ASFV entry. After attachment, ASFV enters the cell via two endocytic pathways: clathrin-mediated endocytosis and micropinocytosis. The fact that ASFV utilizes clathrin-mediated endocytosis to infect host cells is unexpected. This pathway is usually used by small- and intermediate-sized viruses, whose diameter does not proceed 100 nm (Schelhaas, 2010). Nonetheless, detection of ASFV particles within enlarged clathrin-coated vesicles indicates that they can be strained to adapt to larger viruses like ASFV (Hernáez et al., 2016). Other studies showed that ASFV uptake by clathrin-mediated endocytosis requires dynamin GTPase activity, as well as the presence of clathrin-coated pit component Eps15 and cholesterol in cellular membranes (Hernaez and Alonso, 2010; Galindo et al., 2015). Sánchez et al. (2012) used a combination of optical and electron microscopy to show that ASFV utilizes a macropinocytosis-like pathway as the primary means of entry into Vero cells and porcine macrophages. They also showed that ASFV entry was significantly decreased, when cells were treated with inhibitors of the key regulators of macropinocytosis such as Na+/ H+ channels, Pak-1, PI3K, actin cytoskeleton, EGFR, Rac-1 and tyrosine kinases. In addition to monocytes and macrophages, ASFV infects other cells like vascular endothelial cells and lymphocytes. Therefore, it is possible that the usage of macropinocytosis and clathrinmediated endocytosis may increase the ability of ASFV to enter different target cells during the infection process. Once internalized, ASFV particles move throughout the entire endolysosomal system. The inhibitors of endosomal maturation like wortmannin and nocodazole significantly affect ASFV infection, suggesting that virus trafficking throughout endocytic vesicles are essential for a productive infection in target cells (Hernáez et al., 2016). Immediately after infection (15–30 minutes post infection), nearly intact ASFV particles are detectable in early endosomes (Hernáez et al., 2016). At later times (30–90 minutes post infection), viral decapsidation occurs in late endosomes and lysosomes with acidic pH. Interestingly, exposure of purified virus to low pH mimics the observed disassembly of ASFV, indicating that the acidic pH of late endosomes is a signal that triggers ASFV disassembly. Once disassembled, the inner envelope of viral particles fuses with the limiting membrane of the endosomes and naked cores are released into cytoplasm and delivered in perinuclear cytoplasmic viral assembly sites in order to start DNA replication. ASFV factory, gene expression and DNA replication Microtubules provide the major long-range intracellular transport mechanism within the cytoplasm. They play an important role in ASFV transport to the perinuclear area, where viral factories are formed. Early studies showed that incoming ASFV particles were closely associated with microtubules in cytoskeletons obtained Triton X-100 extraction of taxol treated cells (de Matos and Carvalho, 1993). Further studies revealed that the viral structural protein p54 can directly interact with the dynein motor protein and such interaction may constitute a molecular mechanism for microtubule-mediated ASFV transport (Alonso et al., 2001). The viral factory can be characterized as a single and large area localized close to the nucleus, where viral proteins and DNA are accumulated and new virions are assembled. Besides electron microscopy, fluorescent dyes (DAPI or Hoechst) staining cellular and viral

6  | Arabyan et al.

DNA allow the identification of viral factories, where DNA replication occurs. The recombinant ASFV expressing p54 protein fused to the enhanced green fluorescent protein is also used for visualization of ASFV factories (Hernaez et al., 2006). Alternatively, Karalyan et al. (2018) described a simple method of visualizing the viral factory and measuring the number of viral genomes using Feulgen-Naphthol Yellow staining technique and image cytometry. The scheme of viral gene expression revealed that four groups of virus-specific polypeptides, designated as immediate early, early, intermediate and late, are synthesized during ASFV infection in a coordinated manner in which the synthesis of viral DNA presages a major switch in the programme of ASFV gene expression. Immediate early and early genes are expressed before the onset of DNA replication. These genes encode for enzymes that are involved in the nucleotide metabolism and DNA replication, as well as expression of transcription factors that are important for late gene expression (Rodríguez and Salas, 2013). The expression of intermediate and late genes are not detected, when cells are infected in the presence of DNA replication inhibitors, suggesting that their expression depends on the accomplishment of viral DNA replication. These genes encode for structural proteins, polymerases and transcription factors that are packed into the new virions. However, little is known about the mechanisms that control ASFV gene expression during different stages of infection. Recently, in collaboration with Dr Walter Doerfler (Institute for Virology, Friedrich-Alexander-University Erlangen-Nürnberg, Germany), we showed that ASFV DNA does not become de novo methylated in the course of infection, and therefore DNA methylation does not interfere with ASFV gene transcription (Weber et al., 2018). In situ hybridization and radioactive labelling assays demonstrated that ASFV DNA replication has two distinct stages, an early stage in the cell nucleus and a later stage within perinuclear cytoplasmic viral assembly sites (ASFV factory). The genome fragments synthesized in the nucleus are relatively short and not transformed into higher genome length fragments, indicating that the main part of viral DNA replication occurs in the cytoplasm (Rojo et al., 1999). These small DNA fragments exit from the nucleus by disassembling of the lamina network (Ballester et al., 2011). Although the role of nuclear stage in viral DNA replication remains unclear, it has been shown that ASFV growth is inhibited in enucleated cells, suggesting that the nucleus may provide important factors required for viral DNA replication (Ortin and Vińuela, 1977). The precise molecular mechanism by which ASFV DNA replication occurs also remains unclear. However, the presence of head–head and tail– tail genomic intermediates in infected macrophages at late times post infection suggests that ASFV may share a similar replication model to vaccinia virus since the same intermediates have been observed during vaccinia virus DNA replication (Baroudy et al., 1982; Rojo et al., 1999). Data from electron microscopy and in situ hybridization indicate that viral DNA condenses into a pronucleoid structure that is inserted into icosahedral ‘empty’ particles during virion maturation (Brooks et al., 1998). Viral polyproteins pp220 and pp62 are important for the correct assembly and maturation of the core of ASFV particle. Their suppression results in formation and egress of empty viral particles (Suárez et al., 2010). Control of apoptosis and autophagy Infected cells can undergo apoptosis as an antiviral response to viral infection, thereby limiting viral infection. ASFV infection can activate endoplasmic reticulum stress and unfolded protein response. These processes are reflected by activation of caspase-12, leading to the subsequent activation of mitochondrial caspase-9 and effector caspase-13 (Galindo et al.,

African Swine Fever Virus |  7

2012). As ASFV requires a viable cell for replication, the death of the cell limits cellular functions that are required for the virus. Therefore, ASFV encodes for proteins that inhibit cell death, thereby allowing viral infection to continue. Early studies identified two viral genes, A179L and A224L, with similarity to cellular Bcl-2 and IAP family members, respectively (Neilan et al., 1993; Chacón et al., 1995). The A179L protein is conserved among different ASFV isolates and contains domains similar to all BH domains. This protein is expressed at early and late times post infection and localizes at the mitochondria or endoplasmic reticulum (Hernaez et al., 2013). It has been shown that A179L protein can bind to several BH-3 only proteins, including the activated truncated forms of the Bid protein. The binding of A179L to pro-apoptotic Bak and Bax has been also observed (Galindo et al., 2008). Another ASFV protein, A224L, inhibits caspase-3 activation and activates NF-κB (Nogal et al., 2001; Rodríguez et al., 2002). The expression of this protein is detected late during infection and it is incorporated into virus particles. Increased caspase-3 activity has been observed in cells infected with an EP153R gene-deletion mutant, indicating that EP153R protein is a cell-death inhibitor encoded by ASFV. Further studies revealed that EP153R is able to reduce the transactivating activity of the cellular protein p53 following induction of apoptosis (Granja et al., 2004; Hurtado et al., 2004). Autophagy is an important vacuolar process of the cell, leading to lysosomal degradation and recycling of intracellular content, including proteins and organelles. This cellular process can also take over bacterial or viral proteins inside autophagosomes and degrade them directly in autolysosomes. Some virus have developed strategies to escape this degradation, by expression of specific proteins. It has been shown that A179L protein interacts not only with Bcl2 family proteins but also inhibits autophagy by targeting Beclin1 through its BH3 homology domain (Hernaez et al., 2013). The same authors showed that induction of autophagy prior or at the time of infection reduced the number of infected cells, and therefore ASFV inhibited autophagosome formation in cells for productive infection. Pathogenesis The entry of ASFV into pigs occurs via oral or nasal routs, although other routes such as tick bites or scarification are also detected. Depending on circulating viral strain, clinical signs may vary from the highly lethal form with 100% mortality to subclinical and almost unapparent disease. The incubation period of ASF may last for a few days or longer in case of the subacute forms of the disease. However, the primary viraemia occurs until 24 h post infection and virus is detected in almost all organs after 30 h post infection. During viraemia, ASFV is found associated with erythrocytes, neutrophils and lymphocytes (Plowright et al., 1994). It is widely accepted that the destruction of macrophages plays an essential role in the development of pathogenesis, particularly in the impaired haemostasis. Pathological changes in blood cells Early studies showed that ASFV infection is accompanied by leucopenia and thrombocytopenia, but their nature may vary depending on the strain virulence and host factors. During acute infection, pigs demonstrate marked neutropenia and lymphopenia from 2 to 3 days post infection (Karalyan et al., 2012). Interestingly, band-to-segmented neutrophils ratio became ten times higher in infected pigs than in the control group. The large number of other immature cells such as metamyelocytes is also observed during the course of infection

8  | Arabyan et al.

indicating that acute ASFV infection is accompanied by the emergence of immature cells in the host blood (Karalyan et al., 2012). Karalyan and colleagues have also found that more than 10% of blood cells in the final phase of acute infection are represented by atypical lymphocytes (Karalyan et al., 2012a,b). Furthermore, they can be observed in the lymphoid organs, particularly in the spleen and lymph nodes (Zakaryan et al., 2015). These cells have high metabolic activity and altered nuclear shape with hyperdiploid DNA content (Karalyan et al., 2016). However, acute ASFV infection leads to a 2-fold decrease in the number of lymphocytes by the last day of infection (Zakaryan et al., 2015). Affected lymphocyte subsets are macrophages, B-lymphocytes, CD4+ T-helper cells (Ramiro-Ibanez et al., 1997). While previous studies have shown that thrombocytopenia develops at the final stage of acute infection (Anderson, 1986; Anderson et al., 1987), we observed a significant decrease in the number of platelets from the day 3 post infection (Zakaryan et al., 2014), suggesting that the level and extent of pathological changes may vary between different virulent strains. Animals infected with a moderately virulent ASFV strain demonstrate minor changes in the number of circulating blood cells, especially a slight increase in neutrophils and decrease in lymphocytes (Wardley, 1982). In chronically infected pigs, macrophages and B-lymphocytes increase in number by 100% within the first week of infection (RamiroIbanez et al., 1997). However, these values are back to normal after the second week of infection, whereas the number of CD4+ and CD8+ T-lymphocytes increases during the same period of time. The expression of SLA I and SLA II on peripheral mononuclear cells is also elevated, thereby indicating for stimulation of the immune system. At the third week post infection, the expression of SLA I and SLA II is significantly decreased, which may lead to impaired antigen presentation. The recovery of SLA expression during ASFV infection can be associated with an effective immune response against the virus, and thereby with survival of infected pigs. Gross pathological changes Upon infection with a virulent strain, pigs display high fever, severe depression, anorexia, vomiting, watery to bloody diarrhoea, conjunctivitis, accelerated respiratory and pulse rate, abortion in pregnant sows, reddened skin, cyanosis, and incoordination. Acute lethal forms are also accompanied by vascular lesions, which include bleeding in gastrohepatic and renal lymph nodes, petechial haemorrhage, hyperaemic splenomegaly, pulmonary oedema and disseminated intravascular coagulation (Gómez-Villamandos et al., 2003). ASF is classified as a viral haemorrhagic disease due to haemorrhagic lesions consistently reported in organs that do not contain a fixed vascular macrophage population, particularly gastrohepatic and renal lymph nodes. However, haemorrhages also occur in other organs including lung, intestine and heart. Although vascular changes in acute forms of the disease are less intense than in subacute forms of ASF, the final stages of acute infection are characterized by extensive bleeding and changes in coagulation system (Neser and Kotzé, 1987). Initially, it was supposed that endothelial damage was the main cause of bleeding (Colgrove et al., 1969). However, ultrastructure experiments have revealed that when bleeding is observed in the lymph nodes and kidneys, there is no virus replication in the endothelial cells of these organs (Carrasco et al., 1997a). Thus, this hypothesis has been ruled out as the initial cause of haemorrhages, although endothelial damages can be implicated in bleeding in the final stage of infection, when ASFV replication is observed in the capillary endothelium. Phagocytic activation of capillary endothelial cells as an alternative hypothesis has been

African Swine Fever Virus |  9

proposed. Upon ASFV infection, activated endothelial cells coincide with extensive infection and destruction of neighbouring monocytes and macrophages. Therefore, phagocytic activation of endothelial cells is likely due to cytokine release from infected macrophages (Salguero et al., 2005). Pro-inflammatory cytokines such as IL-1 and TNF-α are known to trigger a procoagulant state of the endothelium and activate the coagulation cascade in viral haemorrhagic fevers (Akıncı et al., 2013; Basler, 2017). After the lymph nodes, spleen is the major target organ of ASFV. In acute infection, the spleen becomes six times larger than in healthy pigs, crossing the entire abdominal cavity from one side to the other. Affected spleen has rounded edges and purplish-black colour (Carrasco et al., 1995). Microscopic lesions tend to be more intense in those areas, where macrophages are abundant. Particularly, ASFV replication and cytopathic effect are observed in splenic red pulp macrophages, leading to their massive destruction and activation of the coagulation system, which in turn leads to the accumulation of erythrocytes in splenic cords (Carrasco et al., 1997b). Thus, the main function of the spleen, blood clearance, is impaired during acute ASF. Pigs infected with moderately virulent or low-virulent strains demonstrate mild hyperaemic splenomegaly, which is progressively reversed at the end of infection. Another pathological change is pulmonary oedema, which is observed in pigs infected with highly virulent ASFV strains. The population of macrophages in porcine lungs consists of interstitial macrophages, free rounded alveolar macrophages and pulmonary intravascular macrophages located in pulmonary capillaries adjacent to the endothelium. The latter cells appear to be the main target of ASFV in the lung. The secretory activation of macrophages before the viral entry into the lung indicates that it is induced by circulating chemical mediators produced by primary replication organs (Carrasco et al., 1996, 2002). At the same time, cell debris triggers phagocytic activation of pulmonary intravascular macrophages. This leads to increased intravascular pressure, prompting a number of vascular changes, including interstitial and alveolar oedema. Diagnosis Recognition of the clinical signs of ASF is the first alarm that the virus is circulating among wild or domestic pigs. Early detection allows authorities to rapidly implement control measures in affected areas. However, the diagnosis of ASF is complicated by the fact that other pig diseases, particularly classical swine fever, porcine dermatitis and nephropathy syndrome, swine erysipelas and salmonella have similar clinical signs. Therefore, the rapid, sensitive and specific detection of ASFV is important not only for control measures but also for the differential diagnosis of other pig diseases. According to the World Organization for Animal Health recommendations, ASFV diagnosis includes virus isolation, fluorescent antibody tests, real-time and conventional PCR assays (Oura et al., 2013). For large-scale testing of samples, enzyme-linked immunosorbent assay (ELISA) is also used, although it has lower analytical sensitivity than that of PCR assays (Oura et al., 2013) (Fig. 1.3). Several conventional PCR assays have been described (Steiger et al., 1992; Wilkinson, 2000; Bastos et al., 2003). However, real-time PCR has distinct advantages over conventional PCR, and therefore the latter one is now superseded by real-time PCR assays. King et al. (2003) developed the first real-time TaqMan PCR assay. It targets the viral B646L gene and includes an artificial mimic, thereby validating negative results. This assay was validated

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Figure 1.3  Diagnostic tests available for detection of ASFV in field and laboratory samples.

against 25 different ASFV isolates and it did not cross react with other pig viruses. Zsak et al. (2005) developed an alternative real-time TaqMan PCR assay, which was performed in a single tube containing PCR reagents. This assay also targets the viral B646L gene but it has higher analytical sensitivity than the assay described by King et al. (2003). Another real-time PCR assay was reported by McKillen et al. (2010). This assay was designed against the 9GL gene of the ASFV genome, and was found to be sensitive and specific. It was validated against 15 different ASFV isolates. However, the most sensitive PCR assay was developed based on the Linear-After-The-Exponential-PCR (LATE-PCR) principle. The assay was designed to amplify the B646L gene and was validated using 19 ASFV DNA cell culture virus strains and three tissue samples from infected pigs. The analytical sensitivity is between one and ten copies and it was designed to be used in both laboratory settings and portable PCR machine (Ronish et al., 2011). A single-tube multiplexed real-time PCR assay detecting six different transboundary viruses, including ASFV, was described by Wernike et al. (2013). Other multiplex PCR assays for simultaneous detection and differentiation of porcine viruses were also developed (Haines et al., 2013; Grau et al., 2015; Hu et al., 2016). However, all these assays have one or two log lower analytical sensitivity than other real-time PCR assays. In low-income countries, where authorities cannot afford to purchase expensive PCR systems, the antigen ELISA assay is an attractive alternative for virus detection. Commercially produced antigen ELISA kits (ELISA INGEZIM K3, Ingenasa, Spain; IDvet, France; SVANOVIR, Sweden) are currently available, although very few data on the sensitivity of these assays are known. Vaccines Although different strategies have been employed in order to develop an effective vaccine for this disease, no commercially available vaccine exists to prevent ASF. Approaches used in vaccine development have included live-attenuated ASFV (traditional and engineered), inactivated virus and ASFV subunit vaccines (Zakaryan and Revilla, 2016). During the first outbreaks of ASF in Spain and Portugal, ASFV isolates were collected and used for traditional attenuation by cell passages. Pigs infected with attenuated ASFV develop long-term potent resistance against challenge with homologous but rarely with heterologous strains (Leitão et al., 2001; King et al., 2011; Mulumba-Mfumu et al., 2016). Interestingly, the boundaries of homologous cross-protection are not clear, since closely related ASFV isolates fail to produce cross-protection but diverse ASFV may induce some

African Swine Fever Virus |  11

immunity (King et al., 2011; Lacasta et al., 2015). In addition to this uncertainty, liveattenuated ASFV strains cause several side-effects and post-vaccination reactions including necrotic foci, abortion and pneumonia. Chronic ASF was detected in pigs vaccinated with ASFV isolate attenuated by serial passages in bone marrow cells (Manso-Ribeiro et al., 1963). Hypergammaglobulinaemia and systematic immune activation with increased numbers of B lymphocytes, CD8+ T lymphocytes and macrophages were also observed in vaccinated pigs (Leitão et al., 2001). Thus, the application of live-attenuated ASFV is thwarted by safety issues and poor cross-protection against heterologous strains. Hypothetically, recombinant ASFV strains containing specific deletions of genes are safety alternatives of naturally attenuated ASFV vaccines. In the last two decades comparative genomic research has identified ASFV genes associated with virulence. The deletion mutant ASFV BA71ΔCD2 conferred protection not only against lethal challenge with ASFV BA71 strain but also against the heterologous ASFV E75 (Monteagudo et al., 2017). However, the effect of virulence-related gene deletion on viral attenuation and immunogenicity seems to be strain and genotype dependent. For instance, deletion of the thymidine kinase gene from ASFV strains Malawi and Georgia attenuates these strains but only the TK-deleted Malawi is able to induce a protective immunity in inoculated pigs (Sanford et al., 2016). The deletion of the NL gene from virulent strains attenuates the European E70 strain but has no effect on two other strains (Afonso et al., 1998; Neilan et al., 2002; Reis et al., 2016). Multiple mutations in ASFV strains may produce more attenuated and safer ASFV vaccines but recent studies showed that when two virulence-related genes are deleted from ASFV, the protection of immunized pigs is reduced. For example, it has been shown that the B119L and MGF 360/505 gene cluster-deleted ASFV Georgia strain was incapable of inducing protective immune responses in inoculated pigs (O’Donnell et al., 2016). Interestingly, the same strain containing single deletions in either B119L or the MGF 360/505 gene cluster genes protected animals from challenge with virulent ASFV Georgia strain (O’Donnell et al., 2015a,b). All attempts to induce effective immunity by using inactivated virus have failed. These attempts include purified and inactivated virions, infected glutaraldehyde-fixed macrophages, and detergent-treated, infected alveolar macrophage cell cultures (Forman et al., 1982; Kihm et al., 1987; Mebus et al., 1988). Inactivated ASFV failed to induce protective immunity in pigs even in the presence of modern adjuvants (Blome et al., 2014). The complexity of the virus which contains more than 50 structural proteins may explain this failure. Subunit vaccines containing specific viral antigens and optimized delivery system have lower chances to cause adverse reactions. However, identifying which antigens best stimulate the immune system is a tricky and time-consuming process. It has been shown that pigs co-immunized with p30 and p54 by baculovirus vector conferred protection against lethal challenge with ASFV E75 isolate (Gómez-Puertas et al., 1998), but the combination of p30, p54 and p72 proteins did not protect against lethal challenge with highly virulent ASFV Malawi isolate (Neilan et al., 2004). Similarly, when pigs were immunized with a fusion protein containing CD2v, p30 and p54 coupled to ubiquitin, they were not protected against ASFV E75 isolate (Argilaguet et al., 2013). Recently, Jankovich et al. (2018) delivered 47 antigens to pigs by DNA prime and recombinant vaccinia virus boost and then pigs were challenged with a lethal dose of ASFV Georgia 2007/1 isolate. All pigs developed clinical signs consistent with acute ASFV. However, further studies are required in order to identify effective viral antigens for induction of robust protective immunity in pigs. Improved

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immunization strategies including delivery of the putative ASFV antigens, doses and their proper presentation to the host also need further studies. Antiviral drugs Although it is always better to prevent disease rather than to treat, a possible application of antivirals developed at a reasonable cost may have very beneficial effects. Antiviral therapy can prolong the host survival allowing the infected pigs to generate a productive immune response against ASFV. Therefore, such therapies can be applied in affected farms in order to isolate the epidemic area and prevent further spread. Nucleoside analogues are the major group of antiviral agents that inhibit viral replication by incorporation into viral nucleic acids or interfering with their essential enzymes such as polymerases. Various nucleoside analogues were tested for their antiviral activity against ASFV infection in vitro (Gil-Fernández and De Clercq, 1987; Gil-Fernández et al., 1987; Arzuza et al., 1988). (S)-9-(3-hydroxy-2 phosphonylmethoxypropyl)adenine and carbocyclic 3-deazaadenosine were found to exhibit a much higher anti-ASFV activity than other nucleoside analogues. Recently, we found that two nucleoside analogues, 5-(Perylen3-ylethynyl)-arabino-uridine (aUY11) and 5-(Perylen-3-ylethynyl)uracil-1-acetic acid (cm1UY11), possess a potent, dose-dependent inhibitory effect on ASFV infection in Vero cells and porcine macrophages (Hakobyan et al., 2018). The major antiviral effect (3.5 log reduction) was observed when we added aUY11 and cm1UY11 at the internalization stage of ASFV infection. These compounds belong to a novel family of nucleoside analogues, called RAFI (rigid amphipathic fusion inhibitors) (Fig. 1.4). It has been previously shown that RAFIs inhibit the infectivity of unrelated enveloped viruses by targeting the envelope A

Chemical Formula: C31H22N2O8 Molecular Weight: 518,5250

Chemical Formula: C28H16N2O4 Molecular Weight: 444,4460

B

Figure 1.4  Rigid amphipathic fusion inhibitors. (A) 5-(Perylen-3-ylethynyl)-arabino-uridine (aUY11) and 5-(Perylen-3-ylethynyl)uracil-1-acetic acid (cm1UY11) have antiviral activity against ASFV. (B) Other RAFIs with unknown biological activity.

African Swine Fever Virus |  13

lipids to prevent the curvature changes required for the fusion of viral and cellular membranes during viral entry (St Vincent et al., 2010; Colpitts et al., 2013). Thus, it is possible that aUY11 and cm1UY11 can serve as antiviral agents for interference with ASFV infection because their antiviral effect likely relies on biophysical mechanisms. Because of low side effects and high availabilities, natural compounds have been the centre of attention among researchers working in antiviral drug discovery (Zakaryan et al., 2017). For example, sulfated polysaccharides were shown to affect ASFV attachment and subsequent replication because of negatively charged sulfate groups interacting with positively charged amino acids in the viral envelope (García-Villalón and Gil-Fernández, 1991). The aqueous extracts from different plants and marine microalgae demonstrated anti-ASFV activity in a dose-dependent manner (Fabregas et al., 1999; Fasina et al., 2013). Polyphenolic phytoalexins, such as resveratrol and oxyresveratrol, significantly reduced ASFV production by inhibiting viral DNA replication, late viral protein synthesis and viral factory formation (Galindo et al., 2011). Similar results were observed when we treated ASFV-infected Vero cells with 4′,5,7-trihydroxyflavone, also known as apigenin (Hakobyan et al., 2016). This flavonoid was highly effective at the early stages of infection, reducing the viral yield by more than 99.99%. ASFV-infected cells continuously treated with apigenin did not display a cytopathic effect, and ASFV was not detected, neither by viral antigen ELISA nor by conventional titration. Recently, we also reported that genistein, another flavonoid, affected ASFV infection in Vero cells and porcine macrophages (Arabyan et al., Antiviral Research, submitted). The effect was maximal when it was added to cells at middle phase of infection (8 hpi), disrupting viral DNA replication. We revealed the presence of fragmented ASFV genomes in cells exposed to genistein, suggesting that this molecule may act as an ASFV-topoisomerase II poison. Molecular docking studies showed that genistein may interact with four residues of the ATP-binding site of ASFV-topoisomerase II (Asn-144, Val-146, Gly-147 and Leu-148), showing more binding affinity than ATP4–, emphasizing the idea that this viral enzyme can be a good target for drug development against ASFV. Indeed, siRNA knockdown experiments showed that ASFV-topoisomerase II plays an essential role in viral DNA replication (Freitas et al., 2016). Particularly, a significant decrease in the number of both infected cells and viral factories per cell and in virus yields (up to 2.5 log) was found only in cells transfected with siRNA targeting ASFV-topoisomerase II. In conclusion, ASFV represents a significant pig welfare problem in need of immediate redress. Although so far all efforts to generate effective vaccines have failed, further studies to riddle the details of the interaction of ASFV with host cells will provide new opportunities for vaccine development and antiviral drug discovery. We believe that vaccines alone or in combination with antiviral treatments will reduce or completely eliminate the occurrence of disease. References

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Salas, M.L. (1999). African swine fever virus. In: Encyclopedia of Virology, 2nd ed., Webster, R.G., Granoff, A., eds. (Academic Press, London), pp. 30–38. Salguero, F.J., Sánchez-Cordón, P.J., Núñez, A., Fernández de Marco, M., and Gómez-Villamandos, J.C. (2005). Proinflammatory cytokines induce lymphocyte apoptosis in acute African swine fever infection. J. Comp. Pathol. 132, 289–302. Sánchez Botija, A.C. (1963). Reservórios del virus de la peste porcina africana. Investigación del virus de la P.P.A. en los artrópodos mediante la prueba de la hemadsorción. Bull. Off. Int. Epizoot. 60, 895–899. Sánchez, E.G., Quintas, A., Pérez-Núñez, D., Nogal, M., Barroso, S., Carrascosa, Á.L., and Revilla, Y. (2012). African swine fever virus uses macropinocytosis to enter host cells. PLOS Pathog. 8, e1002754. https:// doi.org/10.1371/journal.ppat.1002754 Sánchez-Torres, C., Gómez-Puertas, P., Gómez-del-Moral, M., Alonso, F., Escribano, J.M., Ezquerra, A., and Domínguez, J. (2003). Expression of porcine CD163 on monocytes/macrophages correlates with permissiveness to African swine fever infection. Arch. Virol. 148, 2307–2323. https://doi.org/10.1007/ s00705-003-0188-4 Sanford, B., Holinka, L.G., O’Donnell, V., Krug, P.W., Carlson, J., Alfano, M., Carrillo, C., Wu, P., Lowe, A., Risatti, G.R., et al. (2016). Deletion of the thymidine kinase gene induces complete attenuation of the Georgia isolate of African swine fever virus. Virus Res. 213, 165–171. Sanz, A., García-Barreno, B., Nogal, M.L., Viñuela, E., and Enjuanes, L. (1985). Monoclonal antibodies specific for African swine fever virus proteins. J. Virol. 54, 199–206. Schelhaas, M. (2010). Come in and take your coat off – how host cells provide endocytosis for virus entry. Cell. Microbiol. 12, 1378–1388. https://doi.org/10.1111/j.1462-5822.2010.01510.x Steiger, Y., Ackermann, M., Mettraux, C., and Kihm, U. (1992). Rapid and biologically safe diagnosis of African swine fever virus infection by using polymerase chain reaction. J. Clin. Microbiol. 30, 1–8. Steyn, D.G. (1932). East African virus disease in pigs. In 18th Report of the Director of Veterinary Services and Animal Industry, Union of South Africa 1, pp. 99–109. St Vincent, M.R., Colpitts, C.C., Ustinov, A.V., Muqadas, M., Joyce, M.A., Barsby, N.L., Epand, R.F., Epand, R.M., Khramyshev, S.A., Valueva, O.A., et al. (2010). Rigid amphipathic fusion inhibitors, small molecule antiviral compounds against enveloped viruses. Proc. Natl. Acad. Sci. U.S.A. 107, 17339–17344. https://doi.org/10.1073/pnas.1010026107 Suárez, C., Salas, M.L., and Rodríguez, J.M. (2010). African swine fever virus polyprotein pp62 is essential for viral core development. J. Virol. 84, 176–187. https://doi.org/10.1128/JVI.01858-09 Wardley, R.C. (1982). Effect of African swine fever on lymphocyte mitogenesis. Immunology 46, 215–220. Weber, S., Hakobyan, A., Zakaryan, H., and Doerfler, W. (2018). Intracellular African swine fever virus DNA remains unmethylated in infected Vero cells. Epigenomics 10, 289–299. https://doi.org/10.2217/epi2017-0131 Wernike, K., Hoffmann, B., and Beer, M. (2013). Single-tube multiplexed molecular detection of endemic porcine viruses in combination with background screening for transboundary diseases. J. Clin. Microbiol. 51, 938–944. https://doi.org/10.1128/JCM.02947-12 Wilkinson, P.J. (1984). The persistence of African swine fever in Africa and the Mediterranean. Prev. Vet. Med. 2, 71–82, Wilkinson, P.J. (2000). African swine fever. In Manual of Standards for Diagnostic Tests and Vaccines, 4th ed. (Office International des Epizooties, Paris), pp. 189–198. Zakaryan, H., Arabyan, E., Oo, A., and Zandi, K. (2017). Flavonoids: Promising natural compounds against viral infections. Arch. Virol. 162, 2539–2551. https://doi.org/10.1007/s00705-017-3417-y Zakaryan, H., Cholakyans, V., Simonyan, L., Misakyan, A., Karalova, E., Chavushyan, A., and Karalyan, Z. (2015). A study of lymphoid organs and serum proinflammatory cytokines in pigs infected with African swine fever virus genotype II. Arch. Virol. 160, 1407–1414. https://doi.org/10.1007/s00705015-2401-7 Zakaryan, H., Karalova, E., Voskanyan, H., Ter-Pogossyan, Z., Nersisyan, N., Hakobyan, A., Saroyan, D., and Karalyan, Z. (2014). Evaluation of hemostaseological status of pigs experimentally infected with African swine fever virus. Vet. Microbiol. 174, 223–228. https://doi.org/10.1016/j.vetmic.2014.08.029 Zakaryan, H., and Revilla, Y. (2016). African swine fever virus: Current state and future perspectives in vaccine and antiviral research. Vet. Microbiol. 185, 15–19. https://doi.org/10.1016/j. vetmic.2016.01.016 Zsak, L., Borca, M.V., Risatti, G.R., Zsak, A., French, R.A., Lu, Z., Kutish, G.F., Neilan, J.G., Callahan, J.D., Nelson, W.M., et al. (2005). Preclinical diagnosis of African swine fever in contact-exposed swine by a real-time PCR assay. J. Clin. Microbiol. 43, 112–119.

Classical Swine Fever Virus Sandra Blome*

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Friedrich-Loeffler-Institut, Institute of Diagnostic Virology, Greifswald-Insel Riems, Germany. *Correspondence: [email protected] https://doi.org/10.21775/9781910190913.02

Abstract Classical swine fever (CSF) remains one of the most important threats to profitable and sustainable pig production world-wide and its occurrence in domestic and wild pigs has to be reported to the World Organization for Animal Health (OIE). The causative agent, CSF virus (CSFV), is a small enveloped RNA virus of the genus Pestivirus in the Flaviviridae virus family. The clinical picture of CSF depends on virus and host factors and is highly variable. It can range from an almost inapparent infection to a haemorrhagic fever-like illness with high mortality. An immunopathogenesis with dysregulated cytokine responses is suggested for many lesions. After implementation of strict control measures, several countries with industrialized pig production succeeded in eradicating CSF. These control measures often included mandatory vaccination with live attenuated vaccines that have proven to be safe and highly efficacious. Nevertheless, in most parts of the world, CSF is at least sporadically present in either domestic pigs or wild boar. Endemicity can be assumed in several countries of South and Central America, parts of Eastern Europe and neighbouring countries, as well as Asia, including India. This chapter summarizes the virus properties, pathogenesis and clinical picture as well as control options. Taxonomy Classical swine fever virus (CSFV) belongs to the genus Pestivirus, a growing group of small enveloped RNA viruses within the Flaviviridae family. The members of the genus affect mainly cloven-hoofed animals, i.e. cattle, goat, sheep, pigs, and sometimes wild ungulates. Under natural conditions, CSFV is rather restricted to wild and domestic pigs although some reports suggest that it can also occur in bovine species. Up to very recently, only four pestivirus species were officially recognized by the International Committee on Taxonomy of Viruses (CSFV, bovine viral diarrhoea virus types 1 and 2, and border disease virus), but the group of so-called atypical pestiviruses has grown along with the description of several novel pestiviruses (Blome et al., 2017a). Among them are viruses that can be found in pigs, e.g. Bungowannah virus (found in Australia; Kirkland et al., 2015), the atypical porcine pestiviruses (rather widespread with detections in several

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countries including the USA, Germany, the Netherlands, Spain, and China; Hause et al., 2015; de Groof et al., 2016; Postel et al., 2016a; Beer et al., 2017; Muñoz-González et al., 2017; Schwarz et al., 2017) and the LINDA virus (Austria; Lamp et al., 2017). Pestiviral sequences were also found in Norwegian rats (Firth et al., 2014) and Rhinolophus affinis bats (Wu et al., 2012). Recently, a new taxonomy of pestiviruses was proposed (Smith et al., 2017). In this case, original species would be re-designated as Pestivirus A (original designation bovine viral diarrhoea virus 1), Pestivirus B (bovine viral diarrhoea virus 2), Pestivirus C (classical swine fever virus) and Pestivirus D (border disease virus). New species would be Pestivirus E (pronghorn pestivirus), Pestivirus F (Bungowannah virus), Pestivirus G (giraffe pestivirus), Pestivirus H (Hobi-like pestivirus), Pestivirus I (Aydin-like pestivirus), Pestivirus J (rat pestivirus) and Pestivirus K (atypical porcine pestivirus). Additional new species for which a full coding sequence is so far missing are discussed. However, for the purpose of this chapter, the species designation will remain classical swine fever virus (CSFV). Morphology and structure The enveloped viral particles consist of four structural proteins, namely the core protein (C), and envelope glycoproteins E1, E2, and Erns (reviewed by, for example, Rümenapf et al., 1991; Thiel et al., 1991; Blome et al., 2017c). Glycoprotein E2 represents the major immunogen and plays an important role for virus attachment. Antibodies against E2 are detected by several diagnostic test systems. It can be found both as homodimer and in complex with E1. Glycoprotein E1 forms the functional fusion complex and is crucial for virus entry through receptor-mediated endocytosis ( Ji et al., 2015). At least one of the cellular receptors is porcine regulatory protein CD46 (Dräger et al., 2015a). Recently, it was speculated for BVDV that CD46 is supported by a second receptor upon entry as CD46 is not internalized (El Omari et al., 2013). The Erns is the second immunogenic envelope protein, inducing antibodies in the host. Reactions towards Erns are used in marker concepts (for both E2 subunit vaccines and chimeric live virus vaccines). Glycoprotein Erns has ribonuclease activity and is usually present in form of disulfide bond homodimers (Schneider et al., 1993). In the initial phase of pestivirus infection, Erns was shown to mediate the initial contact of the virus to the host cells, through its ability to bind to glycosaminoglycans, before E2 binds to the specific cellular receptor. Structural protein Erns is also involved in cell culture adaptation and harbours, like the E2, virulence determinants (Hulst et al., 2000; Tews et al., 2009). The core encloses the positive single-stranded RNA genome of approximately 12.3 kb which is translated into one polyprotein. The virion RNA is infectious and, thus, transfection can be utilized to obtain infectious progeny from viral RNA (Meyer et al., 2015). The open reading frame is flanked by both 3′- and 5′ non-translated regions (NTR). The 3′-terminus is not polyadenylated but carries a short poly-C tract. The 5′-NTR includes the internal ribosomal entry site (IRES) that mediates translation initiation. Co- and post-translational processing of the precursor protein by viral and cellular proteases results in 13 mature proteins, the above-mentioned structural proteins and non-structural proteins Npro, p7, NS2–3, NS2, NS3, NS4A, NS4B, NS5A, and NS5B (Moennig, 1992). The non-structural autoprotease Npro and also the envelope protein Erns are unique to pestiviruses and can be found in all members of the genus. The above mentioned non-structural proteins have various functions in viral replication with NS5B

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acting as RNA-dependent RNA polymerase. NS3 has three enzymatic functions and acts as protease, NTPase and helicase. Moreover, some non-structural proteins interact and modulate the host immune response. This is particularly true for Npro, which blocks host interferon regulatory factor 3 (IRF-3) and thus interferon responses. For flavivirus NS4A and NS4B it is known that they can induce autophagy signalling. It is likely that the same is true for pestiviruses. Viral replication As mentioned above, early steps of viral replication are initiated by Erns- and E2-mediated attachment and receptor binding. Internalization takes place through clathrin-mediated endocytosis. In the cytoplasm, fusion of virus membranes with host endosomal membranes takes place and the genome is released into the cytoplasm. Cap-independent translation occurs at the ribosomes of the rough endoplasmic reticulum (ER); the internal ribosomal entry site (IRES) at the 5` NTR mediates the initiation of translation. During the process of replication, a dsRNA genome is synthesized from the genomic ssRNA. Transcription and replication of the dsRNA provides viral mRNAs and new ssRNA genomes with positive polarity. Virus assembly occurs at the ER and virions bud at the ER before they are transported to the Golgi apparatus. The release of new virions takes place through exocytosis (Moennig, 2008; Ji et al., 2015). Genetic variability and distribution For the purpose of virus characterization and molecular epidemiology, different genomic regions have been sequenced (Dreier et al., 2007). It has to be noted that there is no clear correlation between genotype and virulence and true serotypes are missing. In the past, a 150 nucleotide (nt) fragment of the 5′-NTR and a 190 nt fragment of the E2 encoding region have been most widely used, sometimes supplemented with 409 nt of the polymerase gene NS5B (Paton et al., 2000c). These sequences were collected and made available at the European Union and OIE Reference Laboratory for CSF in Hannover (Greiser-Wilke et al., 2006; Dreier et al., 2007). With the advent of modern sequencing techniques that were also accompanied by a drop in sequencing costs, longer fragments or even whole-genomes have become available and provide a broader picture and deeper insights (Postel et al., 2012, 2016b; Beer et al., 2015). Phylogenetic analyses reveal that CSFV can be divided into three genotypes with three or four subgenotypes and several clusters. Some of these genotypes can be assigned to distinct geographical regions and the data sets were used for molecular epidemiology and adaptation of control measures (Postel et al., 2012). On a global scale, the most prevalent genotype over the past two decades was undoubtedly genotype 2, especially subgenotypes 2.1 and 2.3. These viruses have been repeatedly found in domestic and wild suid populations of Europe and Asia and were often characterized by moderate virulence (Beer et al., 2015). However, field isolates from South and Central America, and the American continent in general, were so far all placed into genotype 1. While strains from Argentina, Brazil, Colombia, and Mexico clustered in subgenotype 1.1, the strains from Honduras and Guatemala were classified as subgenotype 1.3. Cuban isolates were historically placed into subgenotype

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1.2, but were recently determined to form the new subgenotype 1.4 (Postel et al., 2013). Very recently, Ecuadorian and American strains in general have been used to perform phylodynamic studies (Garrido Haro et al., 2018). The study shows that a diversity of genotype 1 strains still predominates. The only exceptions for genotype 1 are Colombian strains (from 2005 and 2006) that belonged to subgenotype 2.2. Ecuadorian and Peruvian strains share high identities and form a cluster (in the referenced paper referred to as subgenotype 1.6). The African continent is almost a black box regarding characterization of occurring CSFV strains. Apart from the subgenotype 2.1 CSFV strain that affected South Africa in 2005 (Sandvik et al., 2005), knowledge about the situation in general and molecular epidemiology specifically is scarce. Little is also known about the CSF situation in the Middle East apart from Israel, where an outbreak of CSF was reported in 2009 that was caused by a subgenotype 2.1 strain (David et al., 2011). On the Asian continent, China shows the highest diversity with strains belonging to almost all 1 and 2 subgenotypes. In Taiwan, genotype 3 is additionally found. In India, the historical subgenotype 1.1 is now accompanied by strains of subgenotypes 2.1 and 2.2 that share high identity with Asian strains, in particular Chinese isolates (reviewed by Beer et al., 2015). Pathogenesis, immune responses and clinical presentation Upon oronasal CSFV infection, primary replication takes place in tonsils and other local lymphoreticular tissues. Subsequently, virus progeny reaches regional lymph nodes via lymphatic vessels and enters the blood circulation (Dunne, 1973). Thereafter, virus is disseminated and reaches spleen, bone marrow, visceral lymph nodes, intestinal lymphatic tissues, and other parenchymal organs. Primary target cells for viral replication are cells of the monocytic lineage, i.e. monocytes, macrophages and also dendritic cells. Furthermore, endothelial and epithelial cells can get infected in the early stages. In the late stages of generalized infection, almost every cell type is susceptible towards CSFV (Lange et al., 2011). The extent and duration of virus distribution is dependent on the clinical course and outcome that are outlined below. Clinical signs usually appear after an incubation period of roughly 1 week (2–10 days) but virus shedding already begins prior to overt disease. Shedding is observed through all se- and excretions (saliva, urine, faeces and semen, nasal and ocular secretions) and lasts until sufficient antibody levels are reached to neutralize and eliminate the virus, or until death (Moennig et al., 2003). The course and outcome of CSFV infection are dependent on several factors on both virus and host side. On the agent’s side, the virulence of the CSFV isolate seems to play the major role and, to a lesser extent, the dose and route of infection. On the host’s side, age and immune status are assumed to play the key role. Genetic variances that may lead to variability in innate antiviral immune responses in different breeds or races were suggested to influence the disease severity as well. However, neither beneficial nor detrimental reaction pattern have been clearly defined to date and are therefore a matter of scientific discussion and research. As a general rule, infection courses can be divided into acute forms and forms with longterm persistence of the virus. While the acute form can result in either recovery or death of the animal (acute-transient or acute-lethal), all forms with long-term persistence seem to

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result in the exhaustion and ultimate death of the affected animal. The forms with persistent replication of the virus include chronic disease upon postnatal infection, persistent infection through immunotolerance upon infection of pregnant sows with virus strains of low or moderate virulence in the susceptible phase of gestation (‘carrier sow syndrome’), and persistent infection of piglets upon very early postnatal challenge (Blome et al., 2017c). All these courses are accompanied by constant shedding of the virus and a lack of antibody detection (the chronic form may result in remittent low-titre antibody detection in the early phase). The mechanisms of both chronic and postnatal persistent infection are far from being understood but seem to follow similar lines. The viral population does not seem to play a major role in this outcome ( Jenckel et al., 2017). In all cases of prolonged or persistent disease, unspecific clinical signs predominate and are accompanied by wasting and secondary infections of the respiratory and gastrointestinal tract. Skin lesions are frequently reported (Fig. 2.1). In general, young immunocompetent animals are most severely affected and serious acute-lethal courses predominate with textbook lesions including haemorrhages and neurological disorders. Young pigs may also show peracute courses with more or less instant death. The clinical signs of acute infection with recent strains (especially of genotype 2) include high fever that lasts for approximately one week in animals that are going to recover and will persist in animals that will succumb to infection. In the first week upon disease onset, lethargy, lack of appetite and huddling will be most frequently seen (Fig. 2.2), sometimes accompanied by different manifestations of conjunctivitis (slight discharge to severe purulent forms; Fig. 2.3). Animals with acute-lethal course will then develop different shades of gastrointestinal and respiratory signs as well as wasting until they reach the final stage with haemorrhagic lesions (Fig. 2.4) and neurological disorders (Fig. 2.5) (Blome et al., 2017c). The outcome after infection with moderately virulent strains is usually decided in the second and third week upon infection (death occurring between days 14 and 25 post infection).

Figure 2.1  Skin lesions in a piglet with postnatal persistence.

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Figure 2.2  Unspecific clinical signs in domestic pigs and wild boar upon infection with a highly virulent CSFV strain (seven days post infection). The animals have high fever (> 40.5°C), they huddle and show reduced feed intake.

Figure 2.3  Conjunctivitis and general depression in a weaner pig upon CSFV infection (highly virulent Peruvian isolate, genotype 1).

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Figure 2.4  Cyanosis and haemorrhagic lesions at the ears and snout. Signs two weeks after infection with a virulent CSFV strain of genotype 1.

Figure 2.5  Moribund animal after infection with a highly virulent CSFV strain (‘Koslov’). The animal showed neurological dysfunctions including staggering gait, convulsions and paresis.

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Most adult pigs show mild, transient disease upon infection with moderately virulent strains that can be influenced by several secondary infections. These disease courses are particularly difficult to diagnose under field conditions. Highly virulent strains may differ with regard to the above mentioned age dependence. These strains, e.g. CSFV ‘Koslov’ and ‘Shimen’, lead to severe and usually lethal courses in all age classes of animals and are for that reason used for vaccine trials as worst-case scenario. Wild boar show quite similar signs upon experimental infection. Under field conditions, animals may be found that show incoordination and lack of timidity towards humans and dogs. When CSFV is introduced into a naïve wild boar population, mortality is usually high and, thus, fallen animals are an important indicator of disease introduction. As mentioned above, secondary or concomitant infections are characteristic for CSF (Moennig et al., 2003). They occur as a consequence of a severe immunosuppression promoted by the depletion of both B- and T-lymphocytes. Immunosuppressive events were associated with the relatively late humoral and cellular immune response which is typical for CSF. In detail, a short-term leukocytosis occurs very early after infection followed by leukopenia, particularly affecting lymphocyte populations (Summerfield et al., 1998, 2001; Sun et al., 2010). T-cell depletion develops rapidly after infection while B-lymphocytes are depleted in later disease stages. Kinetics of T-cell depletion are dependent on the cell subset and the virulence of the isolate. While αβ-T-lymphocytes are generally depleted irrespective of the virulence, γδ-T-cells are mainly reduced after highly virulent infections. However, in advanced stages of infection severe lymphocyte depletion occurs irrespective of the virulence and even develops in absence of severe clinical signs (Petrov et al., 2014). The mechanism behind CSF-mediated lymphocyte depletion is apoptosis (Summerfield et al., 1998). Several proinflammatory cytokines including interleukin (IL) 1α, IL-1β, TNF-α, IL-6 and interferon (IFN)-α were suggested to induce apoptosis in lymphocytes or other leucocytes during CSF. Additionally, redistribution of lymphocytes from peripheral blood to lymphoid tissues due to local inflammatory reactions may also be involved in decreasing peripheral lymphocyte counts. Apoptotic reactions as well as phagocytic and secretory activation can also be observed in several macrophage populations (Goméz-Villamandos et al., 2003). These activated macrophages and their dysregulated cytokine release seem to play a crucial role in pathogenesis while direct damage by the virus could be almost excluded for many lesions occurring in the course of CSFV infection (Goméz-Villamandos et al., 2003). A close correlation was reported between viral infection, inflammation and coagulation dysfunctions including thrombocytopenia. In this respect, CSF pathogenesis seems to resemble that of other viral haemorrhagic fevers. In detail, an overexpression of the proinflammatory cytokines IL-1α, IL-6, and IL-8, along with pro-coagulation factors including tissue factor (TF), vascular endothelial cell growth factor (VEGF), E-selectin and other factors may lead to coagulation dysfunctions by activating platelets and endothelial cells and in addition, increase vascular permeability and vasodilatation. Furthermore dysregulation of IL-10, IL-12, and IFN-γ was suggested to play a role. IL-10 and IFN-γ are among the usual suspects as they were also reported to play a role during filoviral haemorrhagic fevers (Lange et al., 2011). Protective immunity is mediated by both, humoral and cellular-mediated immune response (Suradhat et al., 2007). However, a stronger focus on the cellular level was shown,

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especially concerning mediation of early protection. The cell-mediated immune response is primarily focused on the early and middle stages of disease. This response includes quantitative changes in T-lymphocyte populations comprising cytotoxic T-cells (CTL; CD4– CD8+), T-helper cells (CD4+ CD8–), and mature T-cells (CD3+) along with qualitative changes in cytokine expressions by T-cells (IFN-γ, IL-2, IL-4). With regard to the kinetic of T-cell response, an initial activation of cytotoxic T-cells (CD4– CD8+ cells) and T-helper cells (CD4+ CD8– cells) was shown shortly after infection. During the further disease course, a focus on cytotoxic T-cell response was suggested. The protective effect of T-cells was demonstrated in cases of full protection towards CSFV despite the fact that neutralizing antibodies were absent. An activation of memory T-cells (CD4+ CD8+) was observed during disease course. With regard to the humoral response, quantitative and qualitative changes in B-lymphocytes and immunoglobulins (IgM and IgG) occur upon CSFV infection. In lymphoid organs an early increase of B-cells including plasma cells was detected. In detail, IgM+ cells were shown to rise from day 7 post infection (dpi) and IgG+ cells increase from 11 dpi onwards until virus-specific neutralizing antibodies occur between 10 and 21 days post infection in the peripheral blood. The B-cell increase as well as the differentiation into immunoglobulinproducing plasma cells require stimulatory signals mediated through several cytokines, including IL-4, IFN-γ and IL-2, released by activated monocytes/macrophages and T-cells following CSFV infection (see above). This differentiation mechanism was suggested to be dependent on an increased IL-4 level secreted by T-cells, an eventual predominance of IL-4 over IL-2, and a late decrease of IFN-γ. The relatively late occurrence of neutralizing antibodies may be explained by this late change from cell-mediated to humoral immune response, which is characteristic for CSFV infection. Concordantly to the late development of antibodies, IgG+ cell increase occurs in advanced disease stages until they outnumber the initially predominant IgM+ cells. The protective value of neutralizing antibodies against CSFV infection was demonstrated in vivo and in vitro. CSFV specific antibodies are primarily targeted against the envelope glycoprotein E2 which is known to be the major immunogen of pestiviruses and additionally targeted by cytotoxic T-cells. As mentioned before, the envelope glycoprotein Erns presents an additional antibody target. The same is true for non-structural protein NS2-3. Pathology Pathological lesions found upon CSFV infection are mainly dependent on the course of the infection. Acute-lethal courses are often accompanied by enlarged lymph nodes, haemorrhages and petechiae in many organs, e.g. kidney, urinary bladder, stomach and gall bladder. In some cases necrotizing tonsillitis, diphtheroid necrotizing enterocolitis (Fig. 2.6), lesions in the lymphoreticular system, and non-purulent encephalitis can be observed (Goméz-Villamandos et al., 2006). Splenic infarctions (Fig. 2.7) may occur and are considered pathognomonic for CSF (van Oirschot, 1999a). Severe secondary infection may be present that sometimes obscure the CSF related lesions (Depner et al., 1999). In the chronic form, pathological lesions include atrophy of the thymus, depletion of the lymphoid organs, necrosis and ulceration of the small intestine, colon, and ileocaecal valve (Blome et al., 2017c).

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Figure 2.6  Diphtheroid necrotizing enterocolitis in an animal with postnatal CSFV persistence.

Figure 2.7  Splenic infarctions. These lesions are reported to be pathognomonic for virulent CSFV infection but occur rather seldom with recent CSFV strains.

Diagnostic procedures Rapid and reliable diagnosis is of utmost importance for the timely implementation of control measures against CSF. On the international level, laboratory methods as well as sampling and shipping guidelines can be found in the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals and the EU Diagnostic Manual (European Commission Decision 2002/106/EC). Under routine conditions, CSFV is nowadays diagnosed using well established and validated real-time reverse transcription polymerase chain reaction (RT-qPCR) systems (McGoldrick et al., 1998; Paton et al., 2000a,b; Hoffmann et al., 2005, 2009, 2011; Le Potier et al., 2006; Belák, 2007; Le Dimna et al., 2008; Leifer et al., 2011). Several are also commercially available and easy to perform even by non-expert handlers. Recently, field applicable

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RT-PCRs (Liu et al., 2016) but also alternatives have been designed such as loop-mediated isothermal amplification (LAMP) assays (Chen et al., 2009, 2010; Yin et al., 2010; Zhang et al., 2010a, 2011; Chowdry et al., 2014), primer-probe energy transfer RT-q PCR (Liu et al., 2009b; Zhang et al., 2010b) or insulated isothermal RT-qPCR (Lung et al., 2015). For confirmatory purposes and to obtain virus strains for further characterization, virus isolation is still a standard method that employs different permanent cell lines such as porcine kidney cell lines PK15 or SK6. For rapid confirmation after necropsy, detection of antigen on fixed cryosections of tissues is possible using fluorescence antibody or immuneperoxidase assays (Turner et al., 1968; de Smit et al., 2000b). The available antigen ELISAs are recommended for the use with herd-based testing only. While the sensitivity of panpesti-specific assays (based on the Erns) is usually at least comparable with virus isolation, most CSF specific assays lack sensitivity (Blome et al., 2006). For antibody screening in both domestic pig and wild boar populations different commercially available E2 antibody enzyme-linked immunosorbent assays (ELISAs) are employed. These are accompanied by neutralization assays where confirmation or differentiation is needed. Neutralization assays allow, to a certain extent, differentiation of pestivirus antibodies (Greiser-Wilke et al., 2007). For the use with marker vaccines, reliable DIVA (differentiation of infected and vaccinated animals) assays are needed. Commercially available tests that can accompany both E2 subunit vaccines and chimeric vaccines, target the detection of antibodies directed against glycoprotein Erns (Floegel-Niesmann, 2001, 2003; Blome et al., 2006). In this field, additional diagnostic tests have been developed recently. Due to the increased sensitivity of diagnostic tools (especially RT-qPCR), vaccine virus detections are quite common in oral vaccination campaigns of wild boar and vaccination programmes of domestic pigs. For this reason, different RT-qPCR systems have been developed and tested, these allow differentiation between vaccine and field viruses (genetic DIVA) (Li et al., 2007; Zhao et al., 2008; Huang et al., 2009; Leifer et al., 2009a; Liu et al., 2009a; Widen et al., 2014). Sampling can be the bottleneck of swine fever diagnosis, especially in the case of wild boar, but also in remote areas. For this reason, alternative sampling strategies and sample matrices have been tested for CSF (often combined with African swine fever sampling) especially for wildlife specimens and under rural conditions (Michaud et al., 2007; Prickett and Zimmerman, 2010; Mouchantat et al., 2014; Dietze et al., 2017). However, most of them are not yet in routine use and need further validation (Blome et al., 2017c). Epidemiology Susceptible hosts for CSFV are different members of the Suidae family, particularly domestic pigs (Sus scrofa domesticus) and European wild boar (Sus scrofa scrofa) (Depner et al., 1995; Blacksell et al., 2006). Moreover, the susceptibility of common warthogs (Phacochoerus africanus) and bushpigs (Potamochoerus larvatus) for CSFV was demonstrated (Everett et al., 2011). Classical swine fever virus can be transmitted both horizontally and vertically. Horizontal transmission takes places through direct or indirect contact between infected and susceptible pigs. Important indirect routes include feeding of virus contaminated garbage and mechanical transmission via contact to humans or agricultural and veterinary equipment (van Oirschot, 1999a). Where wild boar are affected, primary outbreaks of domestic

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pigs in the same region were often linked to direct or indirect contact with infected wild boar (Fritzemeier et al., 2000). Upon contact, infection usually occurs through the oronasal route, or less frequently via conjunctiva, mucus membranes, skin abrasions, insemination, and the use of contaminated instruments (de Smit et al., 1999; Moennig and Greiser-Wilke, 2008). Infected pigs show high-titre viraemia and shed virus at least from the beginning of clinical disease until death or specific antibodies have developed (see above). The main excretion routes are by saliva, lacrimal secretions, urine, faeces, and semen. In contrast, chronically infected pigs shed the virus continuously or intermittently until death (van Oirschot, 1999a). Vertical transmission from pregnant sows to their fetuses is possible throughout all stages of gestation and can lead to persistently infected offspring (see carrier sow syndrome). Control measures A binding legal framework exists for the surveillance and control in most countries. Integral parts of the control measures are timely and reliable diagnosis, stamping out of infected herds, establishment of restriction zones, movement restrictions, and tracing of possible contacts. Prophylactic vaccination and other treatments are often also strictly prohibited. However, in Europe, where affected wild boar populations were shown to be an important reservoir for the virus, and acted as a source for reintroduction into the domestic pig population (Fritzemeier et al., 2000; Rossi et al., 2015), emergency vaccination of wild boar has been practised to control the disease (Kaden et al., 2002; von Rüden et al., 2008; Blome et al., 2011; Rossi et al., 2010). Emergency vaccination is also among the options to combat CSF in domestic animals. Furthermore, vaccination is still in use to reduce the disease burden in endemically affected countries. Here, vaccination is often mandatory. Highly efficacious and safe live-attenuated CSF vaccines have existed for decades (van Oirschot, 2003b). The underlying virus strains (e.g. the C-strain of CSFV, the Lapinized Philippines Coronel, the Thiverval or the Japanese guinea pig exaltation negative GPE strain) were attenuated through serial passages in animals (rabbits) or cell culture. These vaccines have been implemented in mandatory control programmes that led, together with strict hygiene measures, to the eradication of CSF from several regions of the world (Greiser-Wilke and Moennig, 2004). At this time, they are still in use in several Asian countries including China (Luo et al., 2014), countries of South and Central America, Trans-Caucasian countries, and Eastern Europe. Vaccines are produced by several manufacturers with different quality management and production systems. The C-strain was also adapted to a bait format for oral immunization of wild boar (Kaden et al., 2000, 2002, 2010) and was recently explored for the vaccination of domestic pigs under backyard conditions (Milicevic et al., 2012; Dietze et al., 2013; Monger et al., 2016). While these vaccines usually have outstanding virtues in terms of onset, spectrum and duration of immunity (Terpstra et al., 1990; Ferrari, 1992; Dahle and Liess, 1995; Kaden and Riebe, 2001; Graham et al., 2012), the main drawback is the lack of a serological marker concept (van Oirschot, 2003b) that would allow differentiation of field virus infected from vaccinated animals (DIVA concept). This is usually less important in endemically affected countries where prophylactic vaccination is carried out to reduce the disease burden and to ensure product safety. In general, there are also no legal obligations to use a certain type of vaccine for an emergency vaccination scenario. However, due to the trade restrictions

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that are imposed on pigs vaccinated with conventional live attenuated vaccines, only DIVA vaccines are considered a feasible option for domestic pigs (Blome et al., 2013). Until very recently, only E2 subunit marker (DIVA) vaccines were available on the market (at present, one E2 marker vaccine is commercially available, Porcilis® Pesti, MSD Animal Health, Unterschleißheim, Germany). These vaccines are safe and were shown to provide clinical protection and limit the spread of CSF (Bouma et al., 1999, 2000; van Oirschot, 1999b; Ahrens et al., 2000; Dewulf et al., 2000; Lipowski et al., 2000; Moormann et al., 2000; de Smit et al., 2000a, 2001; Klinkenberg et al., 2002; van Aarle, 2003). However, they show drawbacks especially in terms of early protection (van Oirschot, 2003a,b) and protection against transplacental transmission (Depner et al., 2001). As a result of these problems, emergency vaccination was hardly implemented in domestic pigs (one exception being Romania). Several research groups have therefore sought to develop a next-generation marker vaccine candidate that would ideally answer all demands with regard to safety, efficacy, DIVA potential, and marketability (Beer et al., 2007). Among the concepts that have been investigated are different vector vaccines based on vaccinia virus, pseudorabies virus or adenoviruses. Other vaccine designs include recombinant attenuated vaccines with chimeric constructs, subunit vaccines based on different expression systems, and RNA/ DNA vaccines (recently reviewed by Blome et al., 2017b). In 2014, the European Medicines Agency (EMA) licensed one of the chimeric marker vaccine candidates, ‘CP7_E2alf ’, after extensive testing in the framework of an EU-funded research project (Reimann et al., 2004; König et al., 2007a,b, 2011; Leifer et al., 2009b; Blome et al., 2012, 2014; Gabriel et al., 2012; Rangelova et al., 2012; Eblé et al., 2013; Renson et al., 2013, 2014; Eblé et al., 2014; Feliziani et al., 2014; Goller et al., 2015; Levai et al., 2015; Dräger et al., 2015b, 2016; Farsang et al., 2017). This new marker vaccine is still under investigation and could be a powerful tool for both emergency vaccination of domestic pigs and also wild boar. Oral emergency vaccination of wild boar with baits has proven to be a potent tool to control the disease in wildlife and to safeguard domestic pigs (Rossi et al., 2015). For this purpose, the above-mentioned C-strain formulations have been used in several European countries including Germany and France. To further optimize the strategy, a DIVA vaccine such as ‘CP7_E2alf ’ could be used. The latter was already tested for use in wild boar under both laboratory and field conditions and could be a medium-term option (König et al., 2007b; Blome et al., 2014; Feliziani et al., 2014). Apart from vaccines, antivirals have been discussed as intervention strategy. The advantage would be the immediate action; drawbacks are economic considerations and consumer concerns. Moreover, loss of action through viral mutations/viral evolution can occur. For CSF, some proof-of-concept data have been obtained for a class of chemical molecules (Vrancken et al., 2008, 2009a,b). However, they were never put into practice. References

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Foot-and-Mouth Disease Virus Francisco Sobrino*, Flavia Caridi, Rodrigo Cañas-Arranz and Miguel Rodríguez-Pulido

3

Centro de Biología Molecular ‘Severo Ochoa’ (CSIC-UAM), Madrid, Spain. *Correspondence: [email protected] https://doi.org/10.21775/9781910190913.03

Abstract Foot-and-mouth disease virus (FMDV) is the prototypic member of the Aphthovirus genus within the Picornaviridae family. This virus causes an acute systemic vesicular disease, footand-mouth disease (FMD), which affects livestock worldwide and causes one of the most feared animal diseases. Here we have addressed different aspects dealing with the biology of this highly variable and transmissible virus that are relevant to understand the viral infectious cycle, including its genome organization, its control of gene expression, the proteins encoded by the FMDV RNA and their known functions, as well as the role they play on cell entry and virus replication and pathogenesis. The characteristics of virus particles and the innate and adaptive responses elicited by this virus are also discussed, as well as current and new strategies for FMD control by vaccination and other antiviral strategies. This chapter also addresses the lesions and clinical signs FMD produce, the current approaches for virus diagnosis and characterization, as well as an overview regarding FMDV control and epidemiology. Molecular biology of FMDV Foot-and-mouth disease (FMD) is an acute and highly contagious viral disease that affects cloven-hoofed animals. The etiologic agent of this disease, FMD virus (FMDV) is a positive-sense, single-stranded RNA virus that belongs to the Aphthovirus genus, Picornaviridae family (for reviews, see Bergmann et al., 2000; Sobrino et al., 2001; Mason et al., 2003; Grubman and Baxt, 2004; Sobrino and Domingo, 2017). The FMDV virion is a 140S particle with a symmetric protein capsid that surrounds a single stranded positive-sense viral RNA molecule of about 8.500 nt in length. This genomic RNA comprises a single open reading (ORF) flanked by two highly structured untranslated regions (UTRs; see Fig. 3.1 and next section) that contain important structural elements involved in viral replication and gene expression (Sáiz et al., 2001; Belsham, 2005). Replication and translation of FMDV RNA occur in the cytoplasm of infected cells, in association with cell membranes (Bienz et al., 1990). FMDV RNA is infectious by itself when transfected into susceptible cells. This feature has allowed the construction of plasmids

44  | Sobrino et al. CODING REGION (SINGLE ORF)

5̓ UTR

IRES S VPg

3̓ UTR

P3

P2

P1

L AUG

SL1

cre

SL2 An

Cn

pseudo-knots

Lab P3

P2

P1-2A Lb 2A VP0

VP3

VP1

2BC2C

3ABC

2C

3AB

3CD

2A VP0

VP3

VP1

VP2

VP3

VP1

2B

VP4

2A

3C

3D

3BBB 2B

structural proteins

2C

3A

3C

3D

non - structural proteins

Figure 3.1  Genomic organization of FMDV.

Fig. 1- Genomic organization of FMDV

encoding the full-length genome sequence (infectious cDNA clones), which are a powerful tool to study the function of different genes and RNA structural motifs (Zibert et al., 1990). The FMDV RNA displays a very high mutation rate because of the virus-encoded RNA polymerase that lacks a proofreading mechanism (Domingo and Holland, 1997). The high mutation rate of FMDV, coupled with its rapid growth and extensive population sizes, result in the rapid evolution of this virus, which contributes to the existence of seven serotypes (A, O, C, Asia1, and those from the South African Territories (SAT) 1, SAT2, and SAT3). In addition, numerous variants and subtypes have been further evolved from each serotype (Domingo, 1990). Genome structure In contrast to most cellular mRNAs, no cap structure is present at the 5′ end of the FMDV RNA, and the 5′ untranslated region (UTR) harbours an internal ribosome entry site (IRES) element that promotes the cap-independent translation initiation of the viral genome. Two in-frame AUG codons, separated by 84 nucleotides, are used to initiate translation of the viral polyprotein, the second one being the most used (Belsham, 1992, 2005; López de Quinto and Martínez-Salas, 1998; Andreev et al., 2007). A small viral-encoded protein, 3B, also known as VPg, is covalently linked to the 5′-terminus (Fig. 3.1). The RNA is polyadenylated at its 3′ end. The viral ORF can be translated into a polyprotein of about 250 kDa, which is subsequently cleaved by two virus-encoded proteinases – Leader (Lpro) and 3Cpro – to yield structural and non-structural proteins (NSPs). The ORF region in FMDV genome can be divided into four functional areas based on the location of the primary cleavages and the functions of mature polypeptides encoded (Grubman and Baxt, 1982) (Fig. 3.1). The L region, which is located at the 5′ end of the capsid component and codes for two alternative forms of Lpro termed Lab and Lb. The P1 region, encoding a precursor for capsid polypeptide, can generate four mature capsid proteins: 1A, AB, AC and 1D, also termed VP4, VP2, VP3 and VP1, respectively. The P2 region encodes three viral proteins (2A, 2B, and 2C) in

Foot-and-Mouth Disease Virus |  45

the middle region of the genome. The P3 region encodes four viral proteins: 3A, 3B, 3Cpro and 3Dpol, among which 3C is a viral protease and 3D an RNA-dependent RNA polymerase. Non-coding RNA elements: the 5′ and the 3′ UTRs It is growing evidenced that structures present at both 5′- and 3′-ends of the viral RNA are required to direct the recognition of the genomes of picornaviruses by the viral RNAdependent RNA polymerase 3Dpol. The FMDV 5′ UTR a long region of about 1300 nt including sequences required for the initiation of viral replication and translation. From the 5′-end of the RNA the following structures are found: (i) The S-fragment that encompasses about 360 nt predicted to form a large hairpin structure (Fig. 3.1), which has been shown to establish RNA/RNA interactions with the viral 3′ end (Serrano et al., 2006) and to bind cellular proteins, poly(rC) binding protein (PCBP) and Poly(A) binding protein (PABP). A role for these interactions in the switch from translation to replication and in the putative circularization of the viral RNA by protein bridges between 5′- and 3′-ends has been proposed (Gamarnik and Andino, 1998). (ii) The poly(C) tract, an almost homopolymeric sequence whose presence is a common feature only with most cardioviruses among the picornaviruses. The poly(C) length varies from about 80 to 400 nt (Harris and Brown, 1977). The role on infectivity of the length of the poly(C) tract is not yet clear. (iii) A pseudo-knots region of about 250 nt in which two to four of these highly structured sequence motifs are predicted, depending on the viral isolate (Clarke et al., 1987; Escarmís et al., 1995) The pseudoknot sequences are precisely deleted/inserted in different viruses, supporting a functional role for these structures that remains unclear. (iv) A ‘cis-acting replication element’ (cre) that includes a conserved motif, AAACA, located within a loop at the end of a stable stem structure (Mason et al., 2002) (Fig. 3.1). The cre is essential for virus replication and is required for in vitro urydylylation of 3B FMDV protein to yield the products VPgpU and/or VPgpUpU that act as primers for viral RNA synthesis (Nayak et al., 2005). In other picornaviruses the cre is present in the RNA coding region which points to differences in the replication strategies among the members of this viral family. (v) The IRES (internal ribosome entry site), present in all picornaviruses, is a complex highly structured element of about 450 nt located downstream the cre and partially overlapping with it. The IRES, present in all picornaviruses, leads the cap-independent translation initiation of viral RNA. The FMDV IRES contains a high degree of secondary and tertiary structure (Belsham and Brangwyn, 1990; Kühn et al., 1990) and has been modelled into a five-domain structure (Pilipenko et al., 1989). The FMDV IRES interacts with a number of cellular proteins, including initiation factors important for normal cellular mRNA translation. A host factor, the polypyrimidine tract binding protein (PTBP), was shown to interact with at least two regions of the IRES (Luz and Beck, 1990; Pilipenko et al., 2000) and deletion of these two sites inhibited both the binding of the protein and in vitro translation (Luz and Beck, 1991). PTBP and another cellular factor, the IRES-specific trans-acting factor (ITAF45), are required for the formation of the 48S translation–initiation complex (Pilipenko et al., 2000; Martínez-Salas et

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al., 2001). Another cell factor, Sam68, was also found to preferentially interact with a UAAA motif within domain 4 (Lawrence et al., 2012; Rai et al., 2015). Additionally, Gem-associated protein 5 (Gemin5) was identified to interact with the IRES of FMDV, inhibiting viral translation (Piñeiro et al., 2013). These RNA/host protein interactions are likely to be key factors in the modulation of the functionality of the FMDV IRES, which can affect, as observed in other picornaviruses (Evans et al., 1985; Skinner et al., 1989), the pathogenicity and virulence of FMDV (Martínez-Salas et al., 1993). (vi) A highly structured region of about 90 nt that is also predicted at the 3′ UTR the FMDV genome, preceding a genetically encoded poly(A) tract. Two conserved stem/loops are predicted in this region that has been shown to interact with the cellular protein PCBP and the 3′-end poly(A) (Rodríguez-Pulido et al., 2007). The 3′ UTR is essential for viral replication (Sáiz et al., 2001), and can mediate long-range interactions with the IRES element as well as with the S hairpin at the 5′-end (Serrano et al., 2006) that are essential for the cap-independent translation of picornavirus RNAs. Viral proteins The IRES-driven translation initiation of the FMDV RNA starts at two AUG codons (Forss et al., 1984) following ribosome recognition of the upstream IRES. The single polyprotein synthesized is sequentially processed to yield first different precursors and finally mature proteins. A primary proteolytic processing renders Lpro, P1-2A, 2BC and P3. The precursors P1-2A, 2BC and P3 are further processed into mature viral proteins and some cleavage intermediates with relative stability, such as VP0, 3AB, and 3CD by 3Cpro, as indicated in Fig. 3.1. Processing of VP0, which is autocatalytic and RNA dependent, generates mature proteins VP2 and VP4. It occurs lately, after capsid assembly and it is necessary for complete maturation of the virion (Arnold et al., 1987; Grubman and Baxt, 2004). As in other picornaviruses, protein precursors may perform functions other than those of their individual components (Gao et al., 2016). The main characteristics and their known implication if the viral cycle of structural and non-structural FMDV proteins are commented below. Leader protein (Lpro) The Lpro is a papain-like cysteine protease (Strebel and Beck, 1986) whose 3D structure has been determined (Guarné et al., 1998). The presence of two alternative initiation codons on the FMDV RNA leads the synthesis in infected cells of two different forms of Lpro (La and Lab) (Clarke et al., 1985), which have been shown to cleave the L/P1 junction within the polyprotein (Fig. 3.1) (Medina et al., 1993). In addition, La and Lab proteases induce the cleavage of the translation initiation factor eIF4G, a component of the cap-binding complex eIF4F that bridges the 5′ capped mRNA to the 40S ribosomal subunit (Devaney et al., 1988), resulting in the inhibition of cap-dependent translation initiation and hence the loss of nearly all host-cell protein synthesis. Whether this cleavage is a direct effect or is mediated through cellular proteases remains to be clarified. In contrast, initiation of FMDV RNA translation only requires the Lpro generated C-terminal eIF4G cleavage product, which binds to the FMDV IRES and interacts with the 40S ribosomal subunit (López de Quinto and Martínez-Salas, 2000; Saleh et al., 2001). The integrity of Lpro is not essential for virus viability since the complete Lb coding sequence can be removed (Piccone et al., 1995). In any case, Lpro is considered as a virulence

Foot-and-Mouth Disease Virus |  47

determinant, likely due to its contribution to shutoff cellular protein synthesis. While leaderless virus replicated at only a slightly lower rate than wild-type virus in cultured cells (Piccone et al., 1995), it was markedly attenuated when injected into cattle and pigs (Mason et al., 1997; Chinsangaram et al., 1998). As discussed in the following sections the inhibition of host-cell protein synthesis by Lpro can contribute to abolishing the ability of the cell to mount an antiviral response, mainly through its ability to block crucial steps in the synthesis of the alpha/beta interferons, a relevant issue for the progression of FMDV infection. Lpro can also induce the cleavage of the cellular protein Gemin5 (Piñeiro et al., 2012), whose direct binding to the 3′-end region FMDV IRES inhibits its activity (Pacheco et al., 2009); this cleavage that has been proposed to enhance the IRES function (Fernandez-Chamorro et al., 2014). Capsid proteins (capsid precursor P1-2A) During the polyprotein processing, excision of 2A from 2B to render the P1-2A precursor is mediated by the 2A, which has been proposed to prevent the formation of the peptide bond at the 2A/2B junction rather than acting as a peptidase that breaks a bond that has been formed (Tulloch et al., 2017). Interestingly, the N-terminus of the P1-2A capsid precursor (see Fig. 3.1) resulting upon L excision contains a motif (GXXXS/T) that allows protein recognition by the cellular myristoylation machinery. This modification of the N-terminus of VP4 is important for the assembly and/or stability of the FMDV and PV capsids (Chow et al., 1987; Abrams et al., 1995). The processing of the FMDV P1-2A precursor to 1AB (VP0), 1C (VP3) and 1D (VP1) is achieved by the 3Cpro (see below) and these products can self-assemble into empty capsid particles (Sáiz et al., 1994; Porta et al., 2013a). The cleavage of VP0 to VP3 and VP1 normally occurs associated with RNA encapsidation, in a manner not yet determined. There is some evidence that VP0 cleavage can occur within assembled empty capsid particles (Curry et al., 1995). The cleavage of the 1D/2A junction by the 3Cpro is the last step in the capsid precursor processing (Ryan et al., 1989). It has been reported that this processing is not essential for capsid assembly or virus infectivity (Gullberg et al., 2013b). The structure and properties of the FMDV capsid, including its ability to interact specifically with cellular receptors, are discussed in the following sections. Proteins from the P2 region The FMDV 2BC precursor is processed to 2B and 2C by the 3C protease (Fig. 3.1). Little is known about the function of these proteins that enclose a hydrophobic transmembrane domain, which suggest their association with cell membranes. Indeed, in PV-infected cells, 2B and 2C are found within membrane-associated viral replication complexes (Bienz et al., 1990). Interestingly, the FMDV 2BC precursor is able to inhibit trafficking of proteins to the cell surface (Moffat et al., 2005) an activity associated with the 3A protein of other picornaviruses (Doedens and Kirkegaard, 1995). In FMDV, neither 3A nor 3B or 3C expressed alone show this activity (Moffat et al., 2007), which has been proposed to account for the reduction of MHC class I surface expression observed in FMDV infected cells (Sanz-Parra et al., 1998). Although the precise role of 2C in RNA replication is not established, single amino acid substitutions within 2C protein can confer resistance to guanidine hydrochloride (Pariente et al., 2003), an inhibitor of the viral RNA replication (Saunders and King, 1982). Recently,

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it has been reported that the 2C protein of human EV71 has both RNA helicase and RNA chaperone activity (Xia et al., 2015). In this line, when expressed in E. coli, FMDV 2C protein can form hexameric structures in vitro in the presence of ATP and RNA (Sweeney et al., 2010), these properties are characteristic of AAA+ ATPases with helicase activity. Proteins from the P3 region The 3A protein The 3A protein is produced by cleavage of 3ABC precursor, and is one of the most variable viral FMDV proteins, being the variable residues preferentially accumulated at its C-terminus (Carrillo et al., 2005). An 18 amino acids long hydrophobic region (HR) is predicted in the middle of the molecule (Moffat et al., 2005; González-Magaldi et al., 2012). In other picornaviruses this hydrophobic domain has been reported to target 3A to intracellular membranes (Choe and Kirkegaard, 2004; Liu et al., 2004) and could contribute to locate the viral replication complex within a membrane context (Datta and Dasgupta, 1994; Fujita et al., 2007). Indeed, it has been proposed that FMDV 3A would interact with membranes through its central HR, exposing its N- and C- termini to the cytosol, where interactions between viral and cellular proteins required for virus replication are expected to occur (González-Magaldi et al., 2014). As part of the 3AB precursor, the 3A protein could also allow the delivery of the 3Bs (VPgs) to the RNA replication complexes. FMDV 3A partially colocalizes with ER and Golgi markers (O’Donnell et al., 2001; García-Briones et al., 2006) and recent evidences point to the involvement of ER exit sites for virus replication, supporting to the involvement of ER in virus replication (Midgley et al., 2013). The 3A protein has been reported to play a role on FMDV host range, as a single amino acid replacement (Q44R) in this protein conferred FMDV the ability to cause vesicular lesions in guinea pigs (Núñez et al., 2001). In addition, deletions and mutations in the C-terminal region associate both to viral attenuation in cattle (Beard and Mason, 2000) and to decreased replication rates in bovine epithelial cells (Pacheco et al., 2003). These mutants remain pathogenic in pigs, which points to the existence of differences in the interaction of 3A with cellular factors between the two species. Brefeldin A is drug that induces fragmentation of the Golgi complex by inhibiting activation of the ADP ribosylation factor Arf1. While PV and other picornaviruses that disrupt Golgi function are extremely sensitive to the effect of brefeldin A (Maynell et al., 1992), FMDV and cardioviruses are rather insensitive to this compound (O’Donnell et al., 2001), supporting the view that diverse cellular factors are required among picornaviruses to recruit cell membranes to form their replication complexes (Martín-Acebes et al., 2008). As reported for PV and coxsackie B virus (Strauss et al., 2003; Wessels et al., 2006), homodimerization of FMDV 3A has been shown to be mediated by polar residues at two α-helices in the N-terminal part of the protein, and this interaction is required for efficient virus replication (González-Magaldi et al., 2012, 2014). The 3B (VPg) protein FMDV 3ABC region shows unique characteristics among picornaviruses, such as encoding three similar, non-identical copies of viral genome-bound 3B protein (Forss and Schaller, 1982; Forss et al., 1984) that serves as a primer for RNA replication (Wimmer, 1982). The

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three copies of 3B are required for both optimal replication in cell culture (Falk et al., 1992) and for virulence in natural hosts (Pacheco et al., 2010). In addition, the C-terminal fragment of FMDV 3A (up to the HR) is considerably longer than those of the other picornaviruses. The FMDV 3B is a small protein (about 11 kDa), and 3B1, 3B2 and 3B3 coding sequences are arranged in tandem in the genome (see Fig. 3.1). A molecule of uridylylated 3B is required for priming of RNA synthesis, becoming linked to the 5′ terminus of both positive- and negative-sense FMDV RNA strands. Uridylylation of 3B requires as a template the cre at the 5′ UTR (Mason et al., 2002; Nayak et al., 2005). Each of the 3Bs has been found attached to genomic RNA (King et al., 1980) and shown to undergo uridylylation in vitro (Pacheco et al., 2003). Deletion of one or more FMDV 3B proteins renders viable viruses that replicate less efficiently than the wt virus, and the single deletion of 3B3 abolished virus viability but this appeared to result from a defect in polyprotein processing rather than a direct effect on RNA replication (Falk et al., 1992). The 3C protease The FMDV 3C protease (3Cpro) is responsible for most of the cleavages within the polyprotein coding sequence. In contrast to the PV 3Cpro (Ypma-Wong et al., 1988), it does not require 3D sequences for its processing activities. FMDV 3Cpro is a member of the chymotrypsin-like family of serine proteases for which the catalytic residues (Grubman et al., 1995) and the 3D structure have been determined (Birtley et al., 2005). FMDV 3C also modifies different cellular proteins as part of the cell subversion mechanism operating during virus infection (see next sections). In addition to its proteolytic activity, FMDV 3C also has RNA binding activity. This probably accounts for the requirement of 3CD (or 3C itself) within the in vitro uridylylation assay (Nayak et al., 2006). The 3D RNA-dependent RNA polymerase, 3Dpol The 3Dpol is a multifunctional enzyme that (i) polymerize nucleotides to synthesize both positive and negative viral RNA strands, and (ii) accomplish the uridylylation of 3B in a process in which the cre motif and 3CD are in vitro required (Nayak et al., 2005). The FMDV RNA polymerase, like those of other RNA polymerases (Ferrer-Orta et al., 2004; FerrerOrta et al., 2006), displays a closed ‘right-hand’ 3D conformation with the fingers, palm and thumb subdomains surrounding different sequence and structural motifs that have been shown to play different roles, reviewed by Ferrer-Orta and Verdaguer (2017). As described for other picornaviruses (Aminev et al., 2003), a nuclear localization signal (NLS) has been identified at the N-terminus of 3D, which has been proposed to mediate the nuclear translocation of 3Dpol and its precursor 3CD in infected cells (Sanchez-Aparicio et al., 2013). This would allow 3Cpro to contribute to the alterations associated to the nuclear reprogramming occurred upon picornavirus infection (Weidman et al., 2003). The combination of low fidelity of replication and the absence of proofreading and excision activities within the 3Dpol result in high mutation frequencies that render quasispecies populations in which a multitude of genomes are subjected to continuous events of mutation, competition and selection (Domingo et al., 1985; Domingo and Schuster, 2016). This quasispecies structure endows FMDV and other RNA viruses for a rapid adaptation to changing environments and antiviral constrains, reviewed by Domingo et al. (2017).

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Capsid morphology and antigenic structure The crystal structure of FMDV particles was one of the first among those of animals viruses to be elucidated in the 1980s (Acharya et al., 1989; Lea et al., 1994). While sharing broad structural similarities with the structure of other members of the Picornaviridae family, FMDV particles exhibited structural differences whose biological relevance has, in several cases, been investigated. The small (30 nm in diameter), roughly spherical FMD virion is formed by a non-enveloped protein icosahedral capsid with a smooth surface. The mature FMDV capsid is made of 60 copies of each of three major structural proteins termed VP1, VP2, VP3 that are exposed in the particle surface, and of the internal, smaller protein VP4 (Figs 3.1 and 3.2; for a current review see Mateu, 2017). VP4 is myristoylated at its N-terminus, and can be regarded as a long N-terminal extension of VP2 that, as discussed above, is released by proteolytic cleavage during virion maturation (Chow et al., 1987). VP1, VP2 and VP3, each about 210–220 residues in length, are structurally similar and display at their core an eight-stranded β-barrel (Acharya et al., 1989). The eight β-strands (termed alphabetically as they appear in the amino acid sequence) form two four-stranded β-sheets (including strands C, H, E, F and B, I, D, G, respectively) and are connected by loops of variable length, each denoted according to the two strands they connect. These surface-exposed loops are involved in the antigenic properties of the virion (Parry et al., 1990; Lea et al., 1994; Curry et al., 1995; Mateu, 2004), as well as on the binding to the cell receptor (Fox et al., 1989). One copy of each of the capsid proteins associate into 60 equivalent, roughly trapezoidal substructures, through non-covalent interactions. This substructure, termed the biological protomer, constitutes the building block from which the capsid is assembled. Five protomers associate around each capsid 5-fold axis to form a higher-order pentagonal capsid substructure termed the pentamer (Fig. 3.2B and C). The protomers in each pentamer are held together by multiple non-covalent interactions weaker than those involved in intraprotomer interactions. The association of the N-termini of VP3 and VP4 around each of the 5-fold axis contributes to connect the protomers in the pentamer. A capsid is made up of twelve pentamers (Fig. 3.2D and E) that are the intermediates of capsid assembly and disassembly.

D

B A

×5 Protomer

×12 Capsid (external view)

Pentamer (external view)

C

Pentamer (internal view)

E

Cápsida

Capsid (internal view)

Figure 3.2  Virus assembly: capsid intermediates. Fig. 2 – Virus assembly: capsid intermediates

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The interface between adjacent pentamers is mainly stabilized by interactions among VP2-VP3 and VP2-VP2 residues at opposites pentamers (Fig. 3.3), held together by electrostatic interactions, hydrogen bridges and weak hydrophobic interactions (Acharya et al., 1989; Lea et al., 1994; Mateo et al., 2003). Unlike those of other picornaviruses, the FMDV capsid is dissociated into 12S pentameric subunits at a pH of below 6.5 (Brown and Cartwright, 1961). It has been proposed that this higher acidic sensitivity is due to the contribution of a cluster of His residues present at the pentamer interface that become protonated at low pH, weakening the capsid stability by electrostatic repulsion (Curry et al., 1995). Antigenic structure The FMDV antigenic sites recognized by B lymphocytes (see next section) are composed of amino acid residues exposed on the surface of the capsid (Acharya et al., 1989). For serotypes A, O C, Asia 1 and SAT 2 A, a major continuous antigenic site (site A) recognized by B lymphocytes is located at the G-H loop, a surface loop connecting the βG and βH strands of capsid protein VP1 (Pfaff et al., 1982; Strohmaier et al., 1982; Bittle et al., 1982; Opperman et al., 2012; Grazioli et al., 2013) (Fig. 3.4). Indeed, a large proportion of monoclonal antibody (MAb) resistant (MAR) mutants obtained using MAbs raised against entire type FMDV particles showed amino acid substitutions within this site (Mateu, 1995). For serotype C, the antigenic structure of the G-H loop is complex, since different overlapping epitopes, defined by their differential ability to react with individual MAbs, have been mapped on it (Mateu et al., 1990). An additional neutralization site has been identified at the C-terminus of VP1 (site C). This site is apparently continuous and independent from the G-H loop in serotypes A and C (Lea et al., 1994). In type O, its vicinity with the G-H loop in the structure of the capsid, as well as competition studies with neutralizing MAbs, suggest that sites A and C conform a single antigenic site composed of discontinuous epitopes (Barnett et al., 1989).

Figure 3.3  Schematic representation of two opposite pentamers in a 2-fold axis of the FMDV capsid. Red lines depict two of the five protomers that build up a pentamer. VP2 y VP3 proteins of adjacent protomers contact the equivalent proteins of the opposite pentamer conforming an intepentameric interface (indicated by an arrow). VP1: green, VP2: magenta, VP3: cyan, and VP4: yellow.

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Figure 3.4  Location of antigenic B cell sites in a FMDV capsid protomer.

Cross-neutralization assays using MAR mutants have allowed the identification of discontinuous antigenic sites, which are not mimicked by lineal peptides, contributed by residues from different capsid proteins in serotypes A and O and Asia 1 (Pfaff et al., 1988; Baxt et al., 1989; Kitson et al., 1990; Grazioli et al., 2013). These antigenic sites are located at exposed regions adjacent to each other and close to the 3-fold axis of symmetry in the capsid. In serotype C, the discontinuous site D, comprises residues involving the C-terminus of VP1, the VP3 B-B knob, and VP2 B-C loop (Lea et al., 1994). Innate and adaptive immune responses FMDV elicits a rapid and broad spectrum of immune mechanisms, including innate responses as well as humoral and cellular adaptive responses (see Fig. 3.4). The combination of these mechanisms results in the induction of efficient protection against infection with homologous and antigenically related viruses, reviewed by McCullough et al. (2017). Innate immune response The innate immune system constitutes the first line of defence in the host cell against pathogen-mediated infections and plays a critical role to induce an antiviral response to suppress viral replication and facilitate the development of specific immune response (Kawai and Akira, 2006; Golde et al., 2008; Summerfield et al., 2009). Innate response is activated through recognition of pathogen-associated molecular patterns (PAMPs) by pattern recognition receptors (PRRs) present in host cells. This recognition activates a downstream signalling cascade culminating in the expression of type-I IFN genes and proinflammatory cytokines that exert antiviral, antiproliferative and immunomodulatory functions (Huber and Farrar, 2011; Tough, 2012). Among the PRRs that are involved in the recognition of FMDV-associated PAMPs are the Toll-like receptors (TLRs), which are expressed on the surface of endosomal and lysosomal membranes of some cell types, including T lymphocytes and antigen presenting cells. The other main PRRs are the retinoic acid-inducible gene

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I (RIG-I)-like receptors (RLRs) (Gay et al., 2014; Lester and Li, 2014), that include RIG-I and the melanoma-differentiation associated gene 5 (MDA5). These molecules are IFNinducible RNA helicases that play an essential role in sensing of viral RNA to triggering the innate immune response (Bruns and Horvath, 2014; Kato et al., 2011). Although RIG-I and MDA5 have similar functions of sensing, differential roles for these two helicases have been suggested (Kato et al., 2006) and MDA5 has been reported as a critical viral sensor for the innate response to picornavirus, including FMDV (Feng et al., 2012). As summarized in Fig. 3.5, binding of viral RNA to the inactivated form of RIG-I or MDA5 initiates innate immunity pathway by activating the mitochondrial antiviral signalling protein (MAVS), through interaction of their ‘caspase activation and recruitment domains’ (CARD) with the corresponding CARD signalling domain of the RLR viral sensor. This interaction triggers a downstream signalling cascade that leads to the activation of the nuclear transcription factor kappa B (NF-kB) and the IFN-regulatory factor 3 and 7 (IRF-3/7), respectively. The nuclear translocation of NF-KB and IRF3/7 stimulate the production of type-I IFN and proinflammatory mediators. The IFN secreted can interact with surrounding host cells by engagement to the IFN-/receptor (IFNAR) on the cell surface, to induce the downstream signalling through the phosphorylation of the tyrosine kinases called ‘Janus kinase’ ( JAKs) and their transcription factors ‘Signal transducers and transcription activators’ (STATs). Once activated, the JAK/STAT complex is translocated to the nucleus where it induces the transcription of hundreds of IFN-stimulated genes (ISGs) to generate an antiviral state in the host aimed at blocking virus replication and dissemination. During co-evolution with their hosts, FMDV has evolved different strategies to avoid innate responses. The FMDV proteases Lpro and 3Cpro besides processing the viral polyprotein to give rise a mature viral proteins (see above sections) are also capable of cleaving .

Figure 3.5  Schematic overview of the antiviral immune response.

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and degrading cellular proteins involved in the innate immune pathway (Galan et al., 2017; Rodríguez-Pulido et al., 2007), thereby enhancing virus replication and suppressing the antiviral response to infection, reviewed in (Tulloch et al., 2017). Indeed, the role of Lpro and 3Cpro on type-I IFN antagonism is being explored intensively, reviewed in (Rodríguez -Pulido and Sáiz, 2017). Analyses of viruses with deletions in the Lpro coding region have demonstrated that FMDV infection is able to block the antiviral activity of IFN-I by limiting the transcription of IFN-I mRNA, inhibiting its translation, as well as those of different ISGs expression (de Los Santos et al., 2006; Zhu et al., 2010). This inhibition was associated to degradation of the p65 subunit of NF-KB by Lbpro and its capability to regulate nuclear activity (de Los Santos et al., 2007). Interferon regulatory factors 3/7 (IRF3/7), two important host proteins for downstream signalling and IFN-I activation, have been also identified as targets of Lbpro. Thus, FMDV Lpro is responsible for decrease of double stranded RNA (dsRNA)-induced type-I IFN production by suppression of IRF3/7 transcription levels (Wang et al., 2010). On the other hand, promoter mutagenesis of the interferon-stimulated response element (ISRE) revealed the importance of FMDV Lbpro to inhibit dsRNA-induced RANTES mRNA transcription by interfering with IRF-3/7 effectors (Wang et al., 2011a). FMDV Lbpro shows remarkable sequence and structural similitudes with viral and cellular deubiquitylation enzymes (DUBs), as well as exhibits a deubiquitinase activity (Wang et al., 2011b). Ubiquitin modification enzymes and DUBs play critical roles in modulating the immune responses (Sun, 2008). Indeed, Lbpro can cleave ubiquitin moieties from some essential proteins of the type-I IFN signalling pathway, such as RIG-I, TRAF family member-associated NF-KB activator (TANK)-binding kinase 1 (TBK1), tumour necrosis factor (TNF) receptor-associated factor 3 (TRAF3), and TRAF6 (Wang et al., 2011b). The interaction of transcription factor host ADNP (activity-dependent neuroprotective protein) and FMDV Lpro has been recently analysed by mass spectrometry (Medina et al., 2017), revealing that Lpro recruits ADNP to the early IFN-α promoter sites during infection and modulates its transcription repression function to decrease IFN and ISGs expression. This interaction suggests a novel role for Lpro to optimize FMDV replication. FMDV 3Cpro can also mediate antagonistic mechanisms to circumvent the innate immune response by cleaving NEMO (IKK-), an essential adaptor for IFR3 and NF-KB activation (Wang et al., 2012), which impairs downstream IFN production. The activity of 3Cpro also antagonizes the JAK-STAT signalling pathway (Du et al., 2014). The blocking of the nuclear translocation of STAT1/2 complex through degradation of karyopherin 1 (KPNA1), the nuclear localization signal receptor for tyrosine-phosphorylated STAT1, by 3Cpro induces a reduction the transcript levels of ISGs and ISRE promoter activity. Several positive-strand RNA viruses, including FMDV, have evolved a novel mechanism to regulate NF-KB signalling by cleavage of TANK (TRAF family member-associated NF-KB activator) (Huang et al., 2015). Although the effects of FMCV 3Cpro remain unclear, it was suggested that cleavage of TANK represent a novel common mechanism adopted for some positive RNA viruses to evade the host innate immune response. In the last years, the effect on viral replication of the expression of individual FMDV proteins, such as 2B, 2C, 3A, VP1 and VP3 has been reported (Li et al., 2013; Zhu et al., 2016), reviewed by Rodríguez-Pulido and Sáiz (2017). Although their mechanisms of action are not well defined, the participation of these proteins in the regulation of the type-I IFN pathway has been proposed.

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In summary, FMDV has evolved to utilize its major proteases to regulate the host antiviral response. Nevertheless, further studies are required to understand the molecular mechanisms involved in the processes of antagonism, as well as their biological relevance. Adaptive immune response Adaptive or acquired immune defence against FMDV has been related to antibody-mediated compartments affording protection in animal models and natural hosts. Induction of specific a response involves B lymphocytes recognizing epitopes on the virus particle to produce specific antibody. Concomitant recognition of T-c cell epitopes following antigen processing and presentation in the context of MHC class II molecules allows stimulation of helper (Th)-lymphocytes to produce growth and differentiation factors necessary for development of the immune response. Critical for inducing effective immune defences are the dendritic cells (DCs) that controls immune defence development and responsiveness, providing essential antigen presentation to T-lymphocytes and antigen delivery to B lymphocytes, reviewed by McCullough et al. (2017). Protection against FMD is often associated with the induction of high levels of neutralizing antibodies in serum (McCullough et al., 1992). However, this response does not warrant clinical protection as animals with low levels of neutralizing antibodies may become protected. It has been proposed that phagocytosis of virus–antibody complexes, following viral opsonization, may mediate viral clearance in vivo (McCullough et al., 1992; Collen, 1994). Neutralizing antibodies directed to B cell epitopes located on the viral capsid (see previous section) can be observed as early as three to four days following infection or vaccination of natural hosts with FMDV, being IgMs those first detected. This response peaks at days 10–14 post infection, and then wanes. IgGs are detected between four and seven days post infection and become the major neutralizing antibodies after two weeks (Francis and Black, 1983). In both, infected and vaccinated animals, IgG1 response is generally greater than that of IgG2 (Mulcahy et al., 1990). Soon after infection or vaccination, there is a detectable antibody response in the secretions of the upper respiratory and gastrointestinal tracts (Francis et al., 1983). The major antibody subclasses found in secretions are initially IgM and then followed by IgA and IgG (reviewed in Salt, 1993). Early evidence of the involvement of antibodies in protection against FMDV drove the attention to the humoral response elicited by the virus. Nevertheless, different findings in the past decades support that T cell responses also contribute to the protective immunity to this virus, reviewed in (Collen, 1994; Sobrino et al., 2001). In cattle and pigs, B cell activation and antibody production are associated with a lymphoproliferative response mediated by T cells (mainly CD4+) that recognize a number of viral epitopes, located in both capsid proteins (Collen et al., 1991; Sáiz et al., 1992; van Lierop et al., 1994; Garcia-Valcarcel et al., 1996; Blanco et al., 2000; Gerner et al., 2009; Liu et al., 2011) and NSP (Rodríguez et al., 1994a; Blanco et al., 2001; García-Briones et al., 2004; Gerner et al., 2006). These T-cell responses mediated by CD4+ cells are expected to be required for protective immunity against FMDV, by participating in the production of antiviral antibodies and by maintaining the appropriate microenvironment needed for a synergistic immune response. FMDVspecific CD8+ T cell responses have also found in conventionally vaccinated pigs and cattle (Sáiz et al., 1992; Guzman et al., 2010). The induction of FMDV-specific effector cytotoxic T lymphocytes (CTL) has been difficult to study and the role CTL activation in protection remains unknown (Rodríguez et al., 1996; Childerstone et al., 1999). On the other hand,

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FMDV infection results in a rapid reduction of MHC class I expression on susceptible cells (Sanz-Parra et al., 1998). Among the consequences of this effect is the impairment of the presentation of viral peptides by FMDV infected cells to CTLs, which may facilitate virus escape from this antiviral mechanism of the host. Virus entry, replication and translation FMDV initiate the infection through the binding to a receptor on the surface of susceptible cells. It is generally accepted that FMDV receptors play a role in tissue and organ tropism that leads to disease pathogenesis (Evans and Almond, 1998; Sobrino et al., 2001). Considering the predilection of FMDV for epithelia, studies were focused on the search for cellular receptor in this cellular type. It is currently accepted that the main physiological cell receptors of FMDV are integrins. These molecules are transmembrane glycoproteins expressed in different cell types that play a role in cell adhesion and migration, thrombosis and lymphocytes interactions. Integrins consist of two subunits (α and β) that are covalently bound (Moreno-Layseca and Streuli, 2014). Binding of FMDV to integrins has been shown to occur through a conserved sequence in VP1 protein, the RGD motif (Robertson et al., 1983; Fox et al., 1989; Parry et al., 1990). This motif is also found in fibronectin, a protein of the extracellular matrix that mediates cell attachment (Pierschbacher and Ruoslahti, 1984). RGD motif is located within the G-H loop that includes amino acid residues 133–158 of VP1 protein (Acharya et al., 1989; Logan et al., 1993) (see Figs 3.4 and 3.6) and is highly exposed and accessible to antibodies, being actually the major B cell antigenic site (site A) of FMDV. Despite the high conservation of the RGD triplet, the G–H loop is highly variable in sequence among FMDVs and it appears that this variation could serve to camouflage these vital residues from antibody attack. Several findings indicate that FMDV may use an alternative receptor present at high copy number for entry into the cell and this receptor is heparan sulfate (HS) (Baxt and Bachrach, 1980; Sekiguchi et al., 1982; Jackson et al., 1996). HS is a glycosaminoglycan (GAG), a polymer of disaccharide repeats that is highly sulfated, and hence negatively charged. HS is widely distributed in animal tissues and among different cell types as an integral membrane component (Kjellén and Lindahl, 1991). FMDV, when adapted to tissue culture, can acquire the ability to use HS by selecting mutations that create a positive capsid surface charge (Baranowski et al., 2000), resulting in the attenuation of virulence in host species (Sa-Carvalho et al., 1997). On the other hand, it has been recently suggested that Jumonji C-domain-containing protein 6 ( JMJD6) might be used as an alternative FMDV receptor in CHO 677 cells (Lawrence et al., 2016). The entry mechanism of FMDV into cultured cells is dependent on the receptor to which the virus binds: integrin-binding FMDV utilizes a clathrin-dependent mechanism (Berryman et al., 2005; O’Donnell et al., 2005; Martín-Acebes et al., 2007); in contrast, HS variants are internalized via the caveola-mediated endocytosis (O’Donnell et al., 2008). A recent work suggests that FMDV can also exploit macropynocitosis to gain entry into the cell (Han et al., 2016) (Fig. 3.6). Penetration and uncoating of FMDV is pH dependent, as shown by using lysosomotropic agents to inhibit viral replication (Carrillo et al., 1985; Baxt, 1987) and by the sensitivity of viral particles to acidic incubation in vitro (Martín-Acebes et al., 2010; Caridi et al., 2015). FMDV uncoating occurs inside early endosomes (Berryman et al., 2005; O’Donnell et al.,

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Figure 3.6  Viral cycle.

2005; Martín-Acebes et al., 2009), where a mild acidic pH induces the breaking down of 140S virion into 12S pentameric subunits and provokes the release of RNA in the cytosol (Cavanagh et al., 1978; Vázquez-Calvo et al., 2014). It has been shown that a genetically engineered FMDV, which is unable to perform the maturation cleavage of VP0 to VP2 and VP4, is non-infectious, can absorb to cultured cells and is acid sensitive (Knipe et al., 1997). This finding suggests that there must be other events after the breakdown of viral particle leading to productive infection. Following uncoating, the RNA is released into the cytoplasm by an as-yet-unknown mechanism, triggering a round of viral translation in an IRES dependent manner. As commented in previous sections, the IRES is a cis-acting RNA element that adopts threedimensional structures to recruit the cellular translational machinery. For most cellular mRNAs, the 5′ cap structure is recognized by a complex composed of eIF4F, eIF4A (an RNA helicase), eIF4E (a cap binding protein) and eIF4G (a scaffold protein), which is recruited to the mRNA along with other factors, resulting in eukaryotic protein synthesis. A specific cleavage of eIF4G by viral Lpro removes its N-terminal portion, the binding sites for eIF4E, impairing cap-dependent protein synthesis in host cells. By contrast, the C-terminal portion of eIF4G retains the binding sites for eIF4A and eIF3, which is sufficient for FMDV IRES activity (Gao et al., 2016).

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Interestingly, the 3′ end of the genome may also be required for FMDV translation, since his deletion rendered non-infectious RNAs with a lowered translation efficiency in in vitro reactions; on the contrary, its addition to a bicistronic construct driven by the FMDV IRES stimulated IRES-directed translation in vitro (López de Quinto et al., 2002). Following initiation, translation results in the production of a single polypeptide, which undergoes a series of cleavages leading to the production of both structural and NS proteins, as discussed in previous sections. How switching from transcription to replication is regulated remains controversial. FMDV 3D and its precursor 3CD localize in the nucleus of cells infected or transiently expressing each of these proteins (García-Briones et al., 2006). Delivery of 3Cpro in the nucleus as part of the 3CD precursor would facilitate the 3C-mediated histone H3 cleavage occurring upon infection (Grigera and Tisminetzky, 1984; Falk et al., 1990). Several additional evidences support the interactions between FMDV proteins and the cell nucleus and the alterations they cause. Thus, RNA helicase A (RHA) (Lawrence and Rieder, 2009) and Sam68, both RNA-binding proteins relevant for viral life cycle, are redistributed from the nucleus to the cytoplasm of infected cells (Lawrence et al., 2012). Also, Lpro is translocated to the nucleus where it mediates degradation of p65/RelA, a subunit of NF-kB (de Los Santos et al., 2007), and of the interferon regulatory factor 3/7 (Wang et al., 2010), as commented in the previous section. Replication of FMDV RNA occurs in the cell cytoplasm in closed association with cell membranes in virus-recruited structures termed replication complexes. The origin of the cell membranes found in these replication complexes remains elusive (Knox et al., 2005). As a RNA polymerase RNA-dependent, 3Dpol first replicates the genomic positive-sense RNA, using the newly generated negative-sense strand as the template for the production of a large amount of new positive-sense infectious genomes (see Fig. 3.6). Viral pathogenesis FMDV produces an acute, systemic vesicular disease, FMD, which affects cloven-hoofed farm animals such as cattle, pigs, sheep and goats, water buffaloes, yaks, llamas, alpacas and Bactrian camels, as well as more than 70 wild ruminant species (Nfon et al., 2017). The basis of this wide host-range remain unknown. In natural infections, the main route of virus entry is the respiratory tract. The virus can also penetrate through skin lesions and it can be experimentally inoculated by intradermal injection into the tongue or in the claws. Despite slight differences between species and the route of inoculation, the initial virus multiplication usually takes place in the pharynx epithelium, producing primary vesicles or ‘aphthae’ (Alexandersen and Mowat, 2005). FMDV proteins are primarily detected within keratinocytes associated to alterations in cells from the stratus spinosum (Fig. 3.7). Within 24–48 h after epithelium infection, fever and viraemia start and the virus enters the blood stream and spreads to different tissues and organs, producing secondary vesicles preferentially in the mouth and feet. The FMD clinical signs include fever, salivating, loss of appetite, lameness and occasionally mastitis (for review see Nfon et al., 2017). Examples of FMD clinical signs and lesions are shown in Fig. 3.7. It has been proposed that macrophages may contribute to the viral spread observed during the viraemia, although the mechanisms behind virus dissemination in infected animals are poorly known (Baxt and Mason, 1995; Yilma, 1980).

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a

d

c

b

e

f

Figure 3.7  Clinical signs and lesions in and around the mouth due to foot-and-mouth disease. (a) Drooling from the mouth. (b) Vesicle on the snout (pig). (c) Fluid-filled elevation and the coronary band extending to the heel bulb (pig). (d) Bovine skin from FMDV experimentally infected bovine showing intraepidermal vesicle (arrow). Note the presence of acantholytic keratinocytes (arrowheads) with neutrophil infiltration. (e) Pig heart showing white streaks and pale areas in the myocardium (arrows). Inset: On cross-section multifocal pale necrotic areas are visible in the left ventricle (arrowheads). (f) Heel of white tailed deer showing immunohistochemical detection of FMDV antigen within keratinocytes (arrow). Adapted from Nfon et al. (2017).

The acute phase of disease lasts about one week and declines gradually, concomitantly with the induction of a strong humoral response. Mortality in adults is relatively low but can be high in young animals associated to an acute viral myocarditis (Fig. 3.7). Secondary bacterial infections in foot vesicles can result in chronic lameness, wasting and mortality. In ruminants, an asymptomatic, persistent infection can be established (van Bekkum et al., 1966). In persistently infected animals the virus can be isolated from the oesophagus and throat fluids from a few weeks up to several years after the initial infection (reviewed in Salt,1993). Both naive and vaccinated animals can become persistently infected (Sutmoller and Gaggero, 1965) following an acute infection. The mechanisms that mediate this persistence are unclear, but it has been proposed that in carrier animals a dynamic equilibrium between the host immune response and the selection of viral antigenic variants at the mucosae of the upper respiratory tract is established (Gebauer et al., 1988). Although still being a controversial issue, there is epidemiological evidence to support that persistently infected animals may be the origin of FMD outbreaks, when brought into contact with susceptible animals (Hedger and Condy, 1985). Diagnosis and viral characterization FMD requires a differential diagnosis with respect to other diseases causing similar clinical signs such as swine vesicular disease (SVD), vesicular stomatitis (VS), vesicular exanthema of swine (VE) and Seneca virus A (SVA). Early recognition and diagnosis of FMD are key

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factors to control disease transmission and are decisive to minimize the consequences of an outbreak, as illustrated in the 2001 epidemic in the UK (McLaws and Ribble, 2007). The diagnostic process usually begins with field inspection by farmers or veterinarians followed by confirmation of the presence of infectious virus or FMDV-specific antibodies in samples using laboratory methods, reviewed in Kitching (2004) and Ludi et al. (2017). The antigenic variation exhibited by FMDV has largely conditioned the strategies followed for its diagnosis, both using genomic and serological procedures. The seven serotypes of FMDV (A, O, C, Asia1, SAT 1–3) have been identified on the basis of the ability of viruses to induce cross-protection in animals This cross-protection is serotype-restricted, and it is not always complete among different subtypes and variants of the same serotype, reviewed in Pereira (1981). Classical techniques, such as complement fixation (CF) – still in use in a limited number of laboratories – and serum neutralization (SN) tests that require the use of established cell lines, have been used for detection of the virus capsid proteins in clinical samples, allowing differentiation between serotypes. ELISA alternatives have also been developed to identify and type FMDV isolates. These ELISAs that usually employ polyclonal antisera in an antigen-capture (sandwich) format and have lower analytical sensitivity to detect FMDV in clinical samples when compared to virus isolation (Ferris and Dawson, 1988). The serological detection of FMDV is carried out by CF, SN and ELISA targeted to detect antibodies against the viral particle (Ludi et al., 2017). This detection is serotype specific and does not allow a reliable distinction between infected and vaccinated animals (Pinto and Garland, 1979). Distinguishing infected from vaccinated animals is important, particularly for cattle, since they can develop a persistent, unapparent infection, even among vaccinated animals (see previous section). Identification of antibodies specific for FMDV infection was formerly performed by detecting the virus infection associated (VIA) antigen, mainly corresponding to the viral protein 3Dpol (Cowan and Graves, 1966). This detection is currently implemented by ELISA assays that detect other NSP other than 3D shown to be specific markers for FMDV infection (Neitzert et al., 1991; Rodríguez et al., 1994a). In particular, ELISAs targeted to 3AB-3ABC antibodies have become a standard procedure for FMDV infection diagnosis (Bergmann et al., 2000; Brocchi et al., 2006). In the last decades, the detection and typing of FMDV RNA by RT-PCR has become a routine test in many laboratories (Meyer et al., 1991; Rodríguez et al., 1994b; Reid et al., 2000) due to its specificity, high sensitivity and the reduced time needed for viral detection. In addition, its combination with direct nucleotide sequencing is an established tool for the rapid characterization of field isolates and the tracing of new outbreaks. Sequencing analyses initially focused on capsid protein VP1 coding sequence (Dopazo et al., 1988; Knowles and Samuel, 2003) are now being extended to the whole genome, reviewed in Ludi et al. (2017). Indeed, the OIE/FAO World Reference Laboratory for foot-and-mouth disease at Pirbright (UK), edits a web page in which an FMDV sequence database is available (http://www. wrlfmd.org/eurl-fmd/). Implementation of extended sequencing to new FMDV isolates is providing relevant information for adequate typing and for vaccine development. FMD remains a serious economic problem for weakest countries of the world, and the technologically more advanced countries should facilitate expertise to control this dreaded disease.

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Foot-and-mouth disease control and epidemiology Despite low mortality rates, FMD severely decreases livestock production and introduces important trade restrictions on animals and livestock products. FMD control in endemic areas is implemented by regular vaccination (see below), which has been instrumental in the eradication of the disease in those areas of the world in which it has been efficiently implemented. Examples are the European Union and, more recently, Uruguay, Argentina, Paraguay and the south of Brazil. A non-vaccination, stamping-out policy (implying slaughtering of infected and contact animals, as well as animal movement restrictions) associated with severe controls in the importation of animals from affected areas was generally adopted to maintaining an area free of FMD since the early 1980s (Donaldson and Doel, 1992). Nevertheless, the increase in global trade favoured in the past two decades the reintroduction of the disease in FMD-free countries with devastating consequences (Sobrino and Domingo, 2001). Examples are the epizootics occurred in 1997 in Taiwan (1997), UK (2001), and Argentina (2000–2001) in which millions of animals had to be slaughtered; in the UK alone more than 10 million cattle, sheep and pigs were slaughtered (reviewed in Rowlands, 2017). The magnitude of these outbreaks and their economic and social costs led to the increasing acceptance of a ‘vaccination to live’ policy as an alternative/complement of the stamping-out policy in case of disease outbreaks in FMD-free countries. This is reflected in the incorporation to the World Organization for Animal Health (OIE) Code of a new category ‘FMD-free country/ zone where vaccination is practised’. Despite reintroductions of FMD in developed countries associated with the increase in global trade, FMD, as many other infectious diseases, is clearly associated with areas with lower levels of development and it contributes to severe economic problems of many developing countries. Control of the disease is hampered by several socio-economic as well as technical factors; among the latter, the considerable antigenic diversity of the virus precludes the preparation of a universal vaccine with unified quality control standards and worldwide distribution. A detailed review on FMD control can be found in Kitching (2004). A huge viral amplification frequently takes place in infected animals, being particularly dramatic in pig, for which up to 1012 infectious units per infected animal have been scored (Sellers, 1971). This makes FMDV to be considered by the OIE as one of the highest transmissible animal viruses. In natural infections, the main route of virus entry is the respiratory tract, and as few as one to ten infective particles can produce the disease (Sellers, 1971). FMDV can be mechanically disseminated by animals, farmers, farming equipment, and during animal transport (Brooksby, 1982). Long-distance, airborne transmission has also been documented (King et al., 1981). Nucleotide sequencing of FMDV RNA and subsequent phylogenetic analyses are increasingly shedding light on the epidemiology of this virus (Martínez et al., 1992). These studies have led to propose that current FMDV lineages (serotypes) probably evolved from a common ancestor infecting African buffalo about 1000 years ago (Knowles, 2013). One of these lineages, SAT serotypes 1–3, remained in Africa in association with African buffalo, the only FMD-susceptible species that can maintain the FMDV infection for long periods of time, probably aided by persistence of SAT infections in a proportion of individuals, and poses an important issue for disease control in Southern Africa, reviewed in (Vosloo and Thomson, 2017). Nucleotide sequencing and phylogenetic analyses are also revealing the patterns of distribution of different serotypes and topotypes, which vary according to

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countries, and in tracing the spatial and temporal spread of different FMD lineages across international boundaries (Knowles et al., 2016). Nucleotide sequencing is also contributing to identify the sources of FMD within outbreaks, a key epidemiological point that can become difficult by limited epidemiological data and the different routes potentially involved in virus spread. Vaccines and antivirals against FMDV Prophylactic vaccination with conventional vaccines has been extensively used to control FMD and has been proven to be the most effective measure for disease control (Parida, 2009). The commercial vaccines currently available are based on chemically inactivated whole viruses that are emulsified with different adjuvants, reviewed in (Smitsaart and Bergmann, 2017). Despite their success in eliciting protective immunity against the disease in endemic countries, these vaccines show several drawbacks (see below) which led to a non-vaccination policy being adopted in the EU in the 1980s. In regions where vaccination is not implemented, massive culling of susceptible animals is the main measure to control the spread of the disease in case of an outbreak, leading to large economic losses due to restriction of animal movement and food industry. The vaccine production was not fully implanted until the middle of the last century when cell culture systems were available and binary ethyleneimine was reported to be an inactivating agent for RNA viruses (Bahnemann, 1972; Mowat and Chapman, 1962). Although, when adequately formulated, these inactivated vaccines induce protective responses, many efforts have been paid to palliate some of their drawbacks, such as (i) the requirement of a constant cold-chain (4°C) to preserve the stability of FMDV particles whose immunogenicity decreases significantly due to is disassociation at higher temperatures into pentamers (see Fig. 3.2), (ii) BSL-3 facilities are needed for production of live viruses, and (iii) the problems associated with ensuring the DIVA (differentiating infected from vaccinated animals) condition of vaccines that has to allow the serological distinction of infected and vaccinated animals (see previous sections). Furthermore, the high variability among FMDVs reflected in seven serotypes and multiple antigenic variants add more complexity to the development of a unique protective vaccine (Taboga et al., 1997). So far, the epidemiology status of each FMD endemic region determines the vaccine formulation including more than one serotype depending on the country. For these reasons, the search of alternative vaccines is a topic of intense research since decades (reviewed in Brown, 1992; Rodriguez and Grubman, 2009; Cao et al., 2016; DiazSan Segundo et al., 2017). Some of those vaccine strategies are commented on below. Live attenuated vaccines The use of non-naturally susceptible animals to produce FMDV vaccines were among the first attempts to obtain attenuated viruses, reviewed in (Bachrach, 1968). This approach implies adaptation of FMDV isolates by serial passages in animal models such us mice or guinea pigs. However, it soon became evident that the use of this type of adapted viruses did not guarantee their attenuation in natural hosts (Núñez et al., 2007). Since then, a balance between attenuation and permissive replication to induce protective immunity has been pursued in the attempts to develop safe live attenuated vaccines. Targeting some of the FMDV virulence factors has been a major strategy to develop life attenuated FMD vaccines.

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In this line, viruses lacking Lpro protein, have been proposed as vaccine candidates. These viruses have the features of being attenuated and inducing neutralizing antibodies in cattle (Mason et al., 1997) and swine (Chinsangaram et al., 1998). Moreover, mutations in the coding region of Lpro render viruses that elicit very early protective immunity (Segundo et al., 2012). The immunogenic potential of FMDVs with mutations in the cell receptor binding motif, RGD, at VP1 capsid protein has also been assessed. Viruses lacking this region were attenuated in cultured cells and in swine, and challenge experiments showed that were able to afford neutralizing antibodies and complete protection in cattle (McKenna et al., 1995). Despite these promising results, the risk of reversion of virulence in natural hosts is perceived as a concern and the major drawback for this type of vaccines. Recent studies reported that engineered viruses with multiple synonymous nucleotide substitutions in the coding region of structural proteins were viable in cell culture while being attenuated in swine (Diaz-San Segundo et al., 2016). This approach exploits the observation that the codon usage bias, a species-specific trait, can influence virus replication and virulence (Mueller et al., 2006). So far, this attenuation approach, called codon deoptimization, has not been tested in FMD challenge experiments. The 3Dpol protein has also been targeted for virus attenuation. Mutations altering the conformation of 3Dpol result in an altered replicative ability in other picornaviruses (Campagnola et al., 2015). In this line, it has recently been shown that mutations affecting fidelity of the RNA polymerase activity led to attenuation in animal models, becoming a potential tool for vaccine development (Rai et al., 2017). Foot-and-mouth disease virus-like empty particles The use of whole empty viral capsids mimicking the intact FMDV particle and lacking infectious RNA has been one of the main areas of research in the past decades, reviewed in Rodriguez and Grubman (2009). The advent of cDNA technology and molecular cloning have made possible the expression of capsid proteins in bacteria and baculoviruses leading to recovery of empty capsids that were immunogenic in natural hosts (Roosien et al., 1990; Grubman et al., 1993). These particles can induce protective immune responses (Xiao et al., 2016) and are considered as DIVA vaccines, since no NSP proteins can contaminate their preparations. Nevertheless, the yield of empty capsids has been shown to be very low, limiting the viability of this approach for vaccine development. As described in previous sections, the efficient expression of FMDV empty viral capsids requires the adequate processing of the P1 polyprotein by 3Cpro. Although the cytotoxic nature of 3Cpro in culture cells has limited this approach for years (Porta et al., 2013b), down-regulation of 3Cpro in the context of a bicistronic system has led to a more stable and efficient production of empty particles (Gullberg et al., 2013a), opening an interesting alternative for the improvement of vaccines based on these particles. Since one of the major drawback for the use empty capsids as vaccines is their thermal stability (empty capsids are more sensitive to heat than virions), attempts are being focused in understanding the molecular basis of this lability and to modify the viral capsids to generate more stable particles (reviewed in, Mateu, 2017). Thus, mutations that introduce disulfide bonds between pentamers led to more thermostable empty- capsids (Porta et al., 2013a). These empty capsids elicited neutralizing antibodies in cattle similar to those of wt capsids and afforded partial protection against challenge (Porta et al., 2013a). Likewise, introduction of mutations in capsid protein VP2, predicted to establish non-covalent interactions that increase pentamer–pentamer stability in type O

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and SAT2 FMDVs, rendered more stable capsids. Calves immunized with these stabilized capsids reached similar neutralizing titres than wt capsids after short-term storage at 4°C. Remarkably, after long-term storage (4–6 months) mutant SAT2-type capsids were able to produce consistent with protection in guinea pigs (Kotecha et al., 2015). In this context, whole-virus inactivated vaccines with improved acid resistance and thermal stability have been reported to be as immunogenic in swine as a commercial vaccine (Caridi et al., 2017). Peptide vaccines Since VP1 was identified as the main target for neutralizing anti-FMDV antibodies, many efforts have been invested in exploring the feasibility of vaccines based on the whole VP1 or fragments of this capsid protein as alternative vaccines. Among their advantages, it is worth highlighting that these vaccines are (i) safe, as a non-infectious material is required, and no reversion to virulence is possible, (ii) DIVA vaccines, (iii) easy to handle and store (no cold chain is required) and (iv) chemically stable. In addition, their large-scale production is affordable. In the early 1980s first attempts to produce synthetic peptides vaccines using VP1-based linear peptides started and Bittle and coworkers reported that a peptide corresponding to the G-H loop in VP1 (see previous sections) induced neutralizing antibodies in mice and protection in guinea pigs (Bittle et al., 1982). Later, it was shown that this peptide, linearly juxtaposed to the C-terminal of VP1, previously reported to induce neutralizing antibodies (Strohmaier et al., 1982), was able to protect cattle from virus challenge (DiMarchi et al., 1986). An important advantage of this B-cell epitope in the VP1 G-H loop (site A) is that is continuous and easy to mimic as a peptide (Wang et al., 2002; Cubillos et al., 2008). In contrast, the discontinuous or conformational-dependent structure of other neutralizing B-cell epitopes (Mateu et al., 1998; Liu et al., 2017) still hamper their mimic using peptides and their use for vaccine purposes (Villén et al., 2002, 2006). Despite the vaccine potential of peptides, the main problem faced during decades has been the weaker immunogenicity they conferred when compared with conventional, inactivated vaccines (Doel et al., 1988). An important consideration regarding VP1-based vaccines is that although the main B-cell epitope at the G-H loop is continuous, it requires a proper folding to become immunogenic, a fact that can has been correlated with the poor protection afforded by bacterially produced VP1 (Kleid et al., 1981). In addition, and relevant for the design of vaccines based on viral subunits, adequate T-cell responses are required to optimize the production of FMDV-neutralizing antibodies and T-cell epitopes have been described in a variety of FMDV hosts, discussed in previous sections and reviewed in (Sobrino et al., 1999). Indeed, inclusion of the sequence of a T-cell epitope described in cattle (Collen et al., 1991) in constructions linearly juxtaposing peptides corresponding to the G-H loop and the site C in VP1 resulted in an improvement in the protection afforded (Taboga et al., 1997). In this large-scale vaccination trial, escape mutants were isolated from unprotected animals, highlighting the potential for virus variation. As classical linear peptides barely achieved levels of protection in livestock compatible with their use as commercial vaccines, multimerization strategies have been developed to overcome their low immunogenicity. Thus, multiple copies in tandem of the VP1 G-H loop from different FMDV serotypes linked to a T-cell epitope have been reported to elicit anti-FMDV responses similar to those of commercial FMD vaccine in mice (Lee et al., 2017). A more complex multimerization approach is the so-called multiple antigenic peptides (MAPs), in which, using a single scaffold molecule, the B-cell epitope is branched

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from a core of lysines resulting in a dendrimer peptide (Tam, 1988; Cubillos et al., 2008). Interestingly, one of this dendrimer peptides displaying four copies of the G-H loop from a type C FMDV linked to a T-cell epitope from FMDV 3A protein, was able to protect pigs against homologous FMDV challenge (Cubillos et al., 2008). Remarkably, downsized versions bearing two copies of the B-cell epitope afforded full protection in swine against an epidemiologically relevant type O FMDV (Blanco et al., 2016), an encouraging result for the validation of these constructs as vaccine alternatives. Antiviral therapies Immunization with conventional vaccines needs a period of about 5–14 days to confer full protection, depending on the vaccine and quality doses. Thus, much attention is being paid to the identification of antiviral strategies to be applied in combination with vaccines to minimize this immunity gap in vaccination schedules. On the other hand, in countries where vaccination is not implemented, antiviral therapies would prevent infection of livestock and susceptible animals in case of sudden outbreaks for which no efficient vaccines were available. The state-of-the-art in FMD antiviral research has been recently and comprehensively reviewed (De Vleeschauwer et al., 2017). Here, we briefly discuss three of the antiviral approaches being currently followed. Biological antivirals approaches Type-I interferons (IFNs) are the main cytokines regulating the innate antiviral response against viral infection including FMDV (Sellers, 1963; Grubman and Baxt, 2004) and, therefore, are candidates to be used as an antiviral tools. Indeed, porcine type-I IFN vectored by human defective-replicative adenovirus (Ad5-pIFN-α) has been reported to protect swine as early as one day post inoculation (Chinsangaram et al., 2003), and this strategy has been extended to type-I IFNs from other FMDV natural hosts (reviewed in Diaz-San Segundo et al., 2017). On the other hand, non-coding (nc) RNAs from the FMDV genome have been reported to protect against infection in in vivo experiments as innate immune response modulators (Rodríguez-Pulido et al., 2011) and the immunogenicity as well as protection conferred by a commercial vaccine is enhanced when these ncRNAs are co-administered in swine (Borrego et al., 2017). Nucleoside analogues The viral RNA-dependent RNA polymerase, 3Dpol for FMDV, is one of the classical targets for pharmacological therapies against RNA viruses (Graci and Cameron, 2008). The first and best-known anti-RNA virus nucleoside analogue ribavirin (1-β-d-ribofuranosyl-1H1,2,4-triazole-3-carboxamide). The inhibition exerted by this compound on the FMDV RNA polymerase activity as well as its potential to induce lethal mutagenesis have been extensively studied (de la Torre et al., 1988; Perales et al., 2011). Favipiravir (T-705) Small molecules have been used as anti-RNA molecules targeting the active sites of different viral proteins. Among them, pyrazinecarboximide derivatives such us T-705 (favipiravir) have been reported to exert a potent antiviral activity against several virus families, including influenza virus (Furuta et al., 2013). It has been reported that cultured cells pretreated

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with T-705 show a reduced susceptibility to FMDV infection, and that this effect was more pronounced when using another pyrazinecarboximide derivative (T-1105) (Sakamoto et al., 2006). Acknowledgement This work was supported by grants AGL2014-52395-C2 and AGL2017-84097-C2-1-R from the Spanish MINECO and P2013/ABI-2906 (co-financed by the Autonomous Community of Madrid and EC FEDER funds). References

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Porcine Circoviruses Sheela Ramamoorthy1*† and Pablo Piñeyro2†

4

1

Department of Veterinary and Microbiological Sciences, North Dakota State University, Fargo, ND, USA. 2 Department of Veterinary Diagnostic and Production Animal Medicine, Iowa State University, Ames, IA, USA. *Correspondence: [email protected] †

Contributed equally

https://doi.org/10.21775/9781910190913.04

Abstract Porcine circoviruses (PCVs), are small, single-stranded DNA viruses belonging to the Circoviridae family. Two major open reading frames (ORF), ORF1 and ORF2, which run in opposing directions on the 1700 bp genome, code for the capsid and replicase proteins, respectively. Transmission of PCV2 occurs both horizontally via contaminated feed, water or contact and vertically from mother to fetus. Porcine circovirus strain 2 (PCV2) was initially identified as the causative agent of post weaning multisystemic wasting syndrome in weanling pigs, which is characterized by severe weight loss and lymphadenopathy. Subsequently, a wide range of clinical syndromes such as reproductive failure, respiratory signs and diarrhoea were associated with PCV2 infections and are now recognized as a part of porcine circovirus-associated diseases or PCVAD. Post-mortem diagnosis of PCVAD requires a combination of the presence of clinical signs and demonstration of PCV2 antigen in tissues or lymph nodes. Ante-mortem diagnosis could involve serological testing for antibodies or detection of viral DNA by PCR in combination with the herd history. Commercial vaccines against PCV2 are effective in preventing clinical signs. They are administered at three weeks of age and prior to farrowing in sows. Both vaccination and strict biosecurity measures are critical for the control of PCV2. However, periodical emergence of new viral strains due to mutation and recombination in the field continue to render PCV2 an economically important pathogen of swine. Introduction Circoviruses are small, non-enveloped, single-stranded DNA viruses belonging to the Circoviridae family. With the recent advances in metagenomic technology, the widespread and ubiquitous nature of circoviruses is being increasingly recognized. Affecting a wide range of mammalian, avian and aquatic hosts, this family is composed of two genera. Circovirus and Gyrovirus, and the proposed genus of Cyclovirus (ICTV, 2016). Porcine circoviruses

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(PCVs), psittacine beak and feather disease virus and other avian circoviruses belong to the Circovirus genus. The pathogenic chicken anaemia virus belongs to the Gyrovirus genus. Human gyroviruses have also been identified but are of unknown pathogenicity. Anelloviruses, such as torque teno viruses, were originally classified under Circoviridae but have recently been assigned a separate family. Cycloviruses infect humans, bats, chimpanzees and dragon flies among other species. Their pathogenicity is unknown; the proposal for their classification as a distinct family is under review (ICTV, 2016). Porcine circoviruses were originally discovered as contaminants of the PK-15 cell line in the 1970s (Tischer et al., 1974), and are widely prevalent in swine populations in several parts of the world (Tischer et al., 1995a,b). However, infection of swine with the contaminated cell cultures did not induce disease. Hence, circoviruses were considered a non-pathogenic part of the host virome (Tischer et al., 1986). In the mid-1990s, increasing number of disease outbreaks in weanling piglets, characterized by severe wasting and lymphadenopathy, led to the discovery of a pathogenic PCV variant. The capsid protein of the new variant differed significantly from the non-pathogenic PCV, both genetically and antigenically (Allan et al., 1998). After Koch’s postulates (Allan et al., 1999) were demonstrated, the newly emerged virus was named porcine circovirus strain 2 (PCV2), while the non-pathogenic variant was called porcine circovirus strain 1 (PCV1). Epidemiology and evolution The initial outbreaks due to pathogenic PCV2 were first documented in Canada (Harding, 2004). The virus spread rapidly within naive swine populations across the world to several countries in Asia, Latin America, Europe and North America. Currently, over 98% of swine herds in the US (Shen et al., 2012), are estimated to be PCV2 positive. This rapid spread can be attributed to animal and animal product travel practices, global trade, virus thermostability, and decontamination difficulties (Meng 2012). Moreover, PCV2 establishes chronic infections and is transmitted in all secretions and excretions (Shibata et al., 2003). Both horizontal and vertical transmission are important in the spread of the virus (Madson et al., 2009). The introduction of commercial vaccines against PCV2 in the US and Europe in 2006 was highly effective in controlling PCV2 and improving production. However, periodical outbreaks were still encountered in vaccinated herds. Investigation of the outbreaks revealed that emergence of a new viral variant, characterized by the presence of a unique motif sequence in the capsid gene. The new variant was named PCV2b, while the virus which emerged earlier in the 1990s was named PCV2a (Ramamoorthy and Meng, 2009). The newly emerged PCV2b variant soon spread rapidly to become the most widely prevalent strain. Thereafter, a third variant, PCV2d, emerged in 2012 and is now the predominant strain in major pork producing regions (Guo et al., 2012). While PCV2a, b and d, have demonstrated virulence properties, PCV2c and e are two other documented genotypes which are not as widely prevalent (Ssemadaali et al., 2015) (Fig. 4.1). More recently, a novel PCV type which is distinct from PCV1 and 2 was found to be strongly associated with the porcine dermatitis and nephropathy syndrome (PDNS) and has been named PCV3 (Palinski et al., 2017). Koch’s postulates for PCV3 are yet to be demonstrated. Plant nanoviruses are considered to be the genetic ancestors of PCV viruses because of similarities in the replicase proteins between the two viruses. A possible recombination event of the plant nanovirus with a RNA virus, and possibly a picornavirus, is believed to had

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Figure 4.1  Phylogenetic tree depicting the pathogenic PCV2 genotypes PCV2a, b and d. The tree was constructed using the MegAlign software, with the neighbour-joining method and the capsid gene sequences. PCV1 was used as the out group. The horizontal distance from the node denotes the extent of genetic differences in sequence between strains.

led to the evolution of PCV1 and the adaptation to mammalian hosts (Gibbs and Weiller 1999). While the exact mechanisms by which PCV2 emerged are unknown, it is clear that PCV2 is continually evolving into new strains; the exact reasons for the rapid evolution are not clear. An overwhelming majority of pigs with clinical signs are infected by more than one genotype of PCV2 (Khaiseb et al., 2011) supporting the theory that recombination is a common mechanism by which viral evolution is achieved. Several recombinant strains containing exchanged genetic segments from the different genotypes are described in literature. The exchanged segments are found in both the ORF1 and 2 genes. Smaller, truncated, replicative PCV2 genomes of about 600–800 bp size have also been detected in pigs. Spontaneous mutations are a second and important mechanism for PCV2 evolution, as the mutation rates for PCV2 are estimated to be 1.2 × 10−3 substitutions/site/year, a rate that is comparatively high for single-stranded DNA viruses (Firth et al., 2009). For example, the recently emerged PCV2d strain and other variants contain one or two additional amino acids in the capsid gene of PCV2b and single amino acid changes in other parts of the ORF2. A clear association of these sequence changes with biological and virulence properties has not been established thus far, probably due to the limitations of experimental models in reproducing clinical signs of the disease (Ssemadaali et al., 2015). Structure and entry Circoviral capsids are non-enveloped, and have the simplest structure for icosahedral viruses with a diameter of about 17 nm. The recent resolution of the crystal structure of the PCV2 capsid protein (Khayat et al., 2011), showed that nucleocapsids contain 60 copies of a single subunit, which assemble in a T = 1 symmetry with 12 vertices, 20 faces and three subunits to each face (Fig. 4.2A). Each subunit contains approximately 234 amino acids, folded into four anti-parallel beta sheets, connected by loops. The beta sheets form the surface topology of the virus, while the loops of one subunit interact with those of the neighbouring subunits to stabilize the icosahedron. The first 40 N-terminal amino acids of the capsid protein contain the nuclear localization signal and are buried within the structure, but could be briefly

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A

B

Figure 4.2  (A) Surface diagram of the PCV2 capsid protein based on the crystal structure, showing the individual subunits arranged in a T=1 symmetry to produce the icosahedral capsid. Reproduced from https://rcsb.org and Khayat et al. (2011). (B) Predicted secondary structure of the individual subunit of 230 amino acids showing the α-helices, β-sheets and flexible regions.

exposed during viral replication (Fig. 4.2B). The capsid protein is immunogenic and the sole component of a majority of the commercially available PCV2 vaccines. Heparan sulfate and chondroitin sulfate B are the putative cellular receptors of the PCV2. Residues 98–103 were predicted to be viral attachment sites as they contained the XBBXBX heparan binding motif (Misinzo et al., 2006). However, with the availability of the crystal structure, these residues were determined be buried within the capsid. Based on the structure, a canyon of positively charged residues located in the 3-fold axis of the capsid is predicted to contain the heparan sulfate binding site (Khayat et al., 2011). The exact mechanisms of viral entry are not known. In epithelial cells, small GTPases and actin polymerization seem to be the mechanisms for viral entry, whereas in monocytes clathrinmediated entry seems to be the mechanism for PCV2 infection. Uncoating of the virus occurs in the low pH of the endolysosome. The mechanisms of nuclear entry and replication are as yet unknown (Misinzo et al., 2006). Genome organization and replication The single-stranded, negative-sense PCV2 genome is approximately 1.7 kb in size. The detection of double-stranded intermediates during replication is not uncommon. While the known information about circoviral genome replication and transcription is derived largely from studies on PCV1, the similarity in the organization of PCV1 and 2 genomes supports the premise that molecular mechanisms would be similar for both viruses. Moreover, the PCV1 and 2 replicase genes are nearly identical and interchangeable, as chimeric viruses replicate well and can infect pigs (Fenaux et al., 2004). The PCV origin of replication consists of a stem–loop structure, flanked by palindromic repeats and situated between the 5′-ends of the two well-characterized open reading frames, ORF1 and ORF2. The full length ORF1 is 936 bp in size and encodes the replicase proteins in a clockwise direction. The ORF2 is 702 (PCV2a and b) or 703 bp (PCV2d) in size, codes for the capsid protein and is anticlockwise in direction (Mankertz et al., 1998). The PCV2 ORF2 gene shares about 60–75%

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homology with the PCV1 ORF2 and about 85–99% homology between PCV2 subtypes. A smaller ORF3 gene is embedded within ORF1 in the reverse direction (Fig. 4.3A). Two major proteins encoded by ORF1 include the Rep and Rep′, which are responsible for binding to the origin of replication initiating replication of the viral genome. Genome replication occurs by the rolling circle melting pot mechanism (Cheung 2004, Cheung 2007), where the replication initiation proteins assemble at the origin to destabilize the secondary structure. The Rep′ protein is believed to have nicking activity and creates a break in the DNA, while the rep protein acts as a helicase to unwind super coils (Fig. 4.3B). Cellular DNA polymerases initiate replication at the free 3′ hydroxyl group generated, to produce one single stranded DNA genome and one double stranded genome for each round of replication. The replicated genomes are either encapsidated or serve as templates for further replication. Transcription Despite the small size of the genome, early studies showed that PCV2 produces at least nine different transcripts during the replication cycle (Cheung, 2003, 2012). With the recent application of next-generation sequencing technology, the number of detected PCV2 transcripts has roughly doubled (Moldován et al., 2017). Two major promoters for the rep and cap genes and other minor promoters drive transcription. The polyA regions of both the cap and rep genes are located in opposing directions in the intergenic region at the 3′ end of the cap and rep genes. An interferon stimulating response element (ISRE), located within the promoter of the rep gene, regulates gene expression in response to host cytokine production during infection (Ramamoorthy et al., 2009). The full-length cap transcript is about 990 nt long and is spliced post-transcriptionally. Several other coding and non-coding RNA rep transcripts have been detected. The major coding transcripts include a full-length rep and two spliced products Rep′ and Rep 3a, b and c. Other non-structural, spliced transcripts called NS0, ND515 and NS672 were detected within the rep coding region. Long noncoding RNAs are RNA species greater than 200 bp in length, which are being increasingly described in several mammalian species and viruses. While the significance of non-coding RNA and the long transcriptss i as yet unknown, two long non-coding transcripts of 2412 and 994 nt length were detected by next-generation sequencing methods for PCV1 (Moldován et al., 2017). These transcripts are named Ctr and Ctr′ (complex transcripts). While little is A

B

Figure 4.3  (A) Genome organization and transcription of PCV2. A. Schematic diagram of the single stranded, DNA genome of ~1.7 kb. P – Promoter, (B) – PCV2 transcripts – Solid lines – Genome or open reading frames, hashed bars – coding segments, dashed lines – spliced segments.

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known about the regulation of viral gene expression, several of the PCV transcripts overlap in a head-to-head or tail-to-tail manner to possibly provide a regulatory function (Fig. 4.3B). The translated products of the rep and cap transcripts result in proteins of 312 and 235 amino acid lengths. The cap and rep proteins are produced during infection and have been recognized in viral cultures using anti-sera from infected animals. The PCV2 capsid protein is necessary and sufficient for inducing vaccine-mediated immunity against PCV2. The ORF3 protein is believed to play a role in PCV2 virulence by regulating apoptosis (Chaiyakul et al., 2010). While earlier predictions and findings regarding the existence of the ORF3 transcript were recently confirmed by next-generation sequencing (Moldován et al., 2017), the ORF3 protein has not been detected in PCV2-infected cells or animals thus far. Clinical presentation The primary disease syndrome associated with PCV2 infections initially was known as the postweaning multisystemic wasting syndrome (PMWS), which was characterized by wasting, rapid loss of condition and lymphadenopathy (Allan et al., 1998). Subsequently, PCV2 has been associated with several other disease syndromes (Segalés et al., 2005). International consensus is still needed regarding a nomenclature that can integrate all of the clinical manifestations associated with PCV2. Collectively, in Europe, the term ‘porcine circovirus diseases (PCVDs)’ is used to denote clinical signs of PCV2 infections. In October 2006, the American Association of Swine Veterinarians (AASV) proposed the name ‘porcine circovirus associated diseases (PCVADs)’ (http://www.aasp.org/aasv/position-PCVADhtm) (accessed 30 November 2014). Although the main focus of research has been systemic PCVAD, also known as PMWS, PCV2 can also be subclinical and has also been implicated as a potential causative agent of porcine dermatitis and nephropathy syndrome (PDNS), porcine respiratory disease complex (PRDC) (Rosell et al., 2000), necrotizing pneumonia (PNP) (Drolet et al., 2003), enteritis (Kim et al., 2004), reproductive failure (Madson et al., 2009; Madson and Opriessnig, 2011) and neuropathy (Stevenson et al., 2001). Subclinical infection The high serological prevalence without clinical presentation of systemic PCVAD (Segalés et al., 2005), and the detection of PCV2 in retrospective studies, even before the first description of PMWS ( Jacobsen et al., 2009), suggest that PCV2 subclinical infection is highly prevalent. Owing to the low prevalence of clinical cases relative to the high prevalence of PCV2 detection, three criteria are necessary for the PCVAD diagnosis: compatible clinical signs, characteristic microscopic lesions and presence of PCV2 within the pathological lesions (Segalés and Domingo 2002). The implications of subclinical PCV2 infections on the health of pigs are as yet uncharacterized. It has been proposed that the efficacy of PCV2 vaccines is impaired in subclinically infected animals (Opriessnig et al., 2006) but conclusive data to substantiate the claim is not available. No detrimental effect in mounting effective immune responses against the pseudorabies vaccine was noted in subclinically infected animals, indicating that chronic PCV2 infection does not interfere with normal host immune responses (Díaz et al., 2012).

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Systemic porcine circovirus-associated diseases (S-PCVAD) S-PCVAD or PMWS was one of the first described and recognized manifestation of PCV2 infection (Ellis et al., 1998). This PCV2 presentation affects mainly grower-finisher pigs from 5 to 16 weeks of age (Gillespie et al., 2009), likely due to the declining levels of maternally derived antibodies (McKeown et al., 2005) at that age. Morbidity varies from 4% to60% and mortality can range from 4% to 20% depending on farm type, husbandry and co-infections. S-PCVAD-affected animals experience weight loss, skin pallor or jaundice, enlarged lymph nodes, dyspnoea and diarrhoea. Less frequent clinical signs also include coughing, pyrexia, and sudden death. Gross pathological changes commonly include enlarged superficial lymph nodes, non-collapsed lungs, gastric ulcers, colonic oedema, and white spotted kidneys (Allan et al., 1999; Rosell et al., 1999; Segalés et al., 2004) (Fig. 4.4). Lymph nodes present a marked follicular and paracortical lymphoid depletion associated with infiltration of histiocytes and/or multinucleated giant cells. Lymphoid depletion associated with granulomatous inflammation can be seen in most of the lymphoid tissues; including Peyer patches, spleen, thymus, and tonsils (Segalés et al., 2004). Histiocytes in granulomatous lesions can also have basophilic intracytoplasmic botryoid inclusion bodies (Rosell et al., 1999). It has been described that severity of the lymphoid depletion is correlated with the amount of PCV2 detected in affected tissues (Darwich et al., 2002). Lymphoid depletion caused by PCV2 has been linked to lymphopenia, specifically depletion of peripheral CD8+ and CD4+CD8++ T and B lymphocytes (Shibahara et al., 2000; Darwich et al., 2002). Other histological changes include multifocal to coalescing lymphoplasmacytic interstitial nephritis, lymphoplasmacytic periportal hepatitis and interstitial pneumonia (Rosell et al., 1999). Depending on the lesion’s chronicity, PCV2 antigen or nucleic acid can be detected by immunohistochemistry (IHC) or in situ hybridization (IHS) in affected tissues (Darwich et al., 2004; Opriessnig et al., 2007). A definitive diagnosis of S-PCVAD requires the detection of the described histological changes and lymph node enlargement, in association with clinical signs (Fig. 4.5).

Figure 4.4  (A) Pigs showing clinical systemic PCVAD signs (post-weaning multisystemic wasting syndrome-PMWS) characterized by loss of weight and thriftiness (Credit College of Veterinary Medicine, Universidad Nacional de la Plata). (B) Pigs showing the characteristic skin lesions of porcine dermatitis and nephropathy syndrome (Credit Iowa State University, Veterinary Diagnostic Laboratory).

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Figure 4.5  (A) Lymphadenopathy in pigs with clinical systemic PCVAD (post-weaning multisystemic wasting syndrome-PMWS). Marked enlarge of inguinal lymph node. (Credit College of Veterinary Medicine, Universidad Nacional de la Plata). (B) Microscopic lesions include severe granulomata lymphadenitis. There is severe lymphoid depletion associated with marked infiltration of histiocytes and multinucleated giant cells. (Credit College of Veterinary Medicine, Universidad Nacional de la Plata).

S-PCVAD or PMWS is considered to be a multifactorial disease, associated with high PCV2 replication levels. However, metagenomic evaluation demonstrated that PCV2 was present equally in PMWS clinically affected and normal pigs, in association with multiple DNA and RNA viruses (Blomström et al., 2009, 2016). However, viral load or detected genetic material was higher in PMWS affected pigs in than healthy pigs, supporting the notion that high viral replication rates are necessary to induce clinical disease (Blomström et al., 2016). Porcine dermatitis and nephropathy syndrome (PDNS) PDNS primarily affects grower and finisher pigs but can also affect adult animals. The prevalence of PDNS is 1–2%, but the mortality is higher, reaching 100% in pigs more than 3 months old and 40–50% in growers (Segalés et al., 1998). Clinical signs of PDNS are non-specific; anorexia, weight loss, depression, and normal temperature to mild pyrexia. The onset of the clinical signs is characterized by animals presenting well demarcated to coalescing red-to-purple macules and papules located on the hind limbs and perineal region. As the disease progresses, lesions have a generalized distribution of dark-red depressed centres covered by serocellular crusts that fade in 2–3 weeks if animals survive the disease, leaving cutaneous scars (Drolet et al., 1999). The kidneys are normally enlarged and have cortical petechiae. Other gross abnormalities include subcutaneous oedema and enlarged and haemorrhagic lymph nodes. Histological lesions are characterized by systemic fibrinonecrotic vasculitis, necrotizing glomerulitis and interstitial nephritis with occasional intratubular proteinaceous casts (Wellenberg et al., 2004). Glomerular sclerosis, interstitial fibrosis and tubular atrophy can also be seen in chronically affected animals. Skin lesions are characterized by dermal and epidermal necrosis associated with vascular necrosis, thrombosis and lymphoplasmacytic perivasculitis (Drolet et al., 1999). While the mechanisms of pathogenesis of PDNS are not fully known, PCV2 seems to be necessary but not sufficient for inducing PDNS manifestation (Rosell et al., 2000).

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Moreover, it has been demonstrated that the viral load of PCV2 is not a decisive factor for the manifestation of PDNS. Previous studies showed animals with PDNS or mild lesions of systemic-PCVAD had no difference in PCV2 viral load (Olvera et al., 2004). The presence of immunoglobulin and complement factors in affected vessels and glomeruli suggest a typical type III hypersensitivity reaction. It has been observed that kidneys from PDNS-affected animals have increased levels of IgG1 + IgG2 and IgM, complement factors C1q and C3, and increased numbers of CD8+ T lymphocytes (Wellenberg et al., 2004). Multiple cofactors associated with the presence of PCV2 have been postulated as causative agents of PDNS. In naturally infected animals, PCV2 and PRRSV, have been detected and isolated from affected tissues (Choi and Chae 2001). Numerous other pathogens such as Staphylococcus hyicus, Actinobacillus pleuropneumoniae, Pseudomonas spp. and Pasteurella multocida (Thomson et al., 2001; Lainson et al., 2002) have also been isolated from tissues with PDNS concomitant with the presence of PCV2. Hence, the exact role of PCV2 in the development of PDNS is still under debate. Recent studies demonstrate that lesions resembling PDNS can be reproduced with PRRSV and Torque Teno Virus (TTV) without the presence of PCV2 (Krakowka et al., 2008). A newly recognized PCV3 has been detected in association with PCV2 in PDNS renal lesions (Palinski et al., 2017). In summary, while a majority of the evidence points to PCV2 as a predisposing factor for PDNS, mechanistic evidence for the role of PCV2 in the pathogenesis of PDNS is currently lacking. Respiratory porcine circovirus-associated disease (R-PCVAD) Porcine respiratory diseases complex (PRDC) is a multifactorial entity that affects pigs from 8 to 22 weeks of age (Opriessnig et al., 2007). Numerous agents have been proposed as causative agents, such as PCV2, swine influenza virus (SIV), PRRSV, pseudorabies virus (PRV), porcine respiratory coronavirus (PRCV), Mycoplasma hyopneumoniae, Pasteurella multocida, Streptococcus suis, and Actinobacillus pleuropneumoniae (Harms et al., 2002). However, it is considered that PCV2 may play an important role, not only as a primary pathogen, but also predisposing to secondary infections (Kim et al., 2003). Perhaps PRRSV, Mycoplasma hyopneumoniae and PCV2 are still the most important pathogens in the PRDC, which is the most common coinfection observed in the field. Regardless of PCV2 genotype, the presence of PRRSV enhance PCV2-associated lesions, increases viral load in serum, and the amount of antigen in tissue (Rovira et al., 2002; Park et al., 2013; Chae, 2016). PCV2 lesions and viraemia are also enhanced by Mycoplasma hyopneumoniae coinfection. However experimental studies demonstrate that pulmonary lesions are more severe after sequential infection compared with simultaneous coinfection (Chae, 2016). Clinical signs are characterized by dyspnoea, cough, lethargy, anorexia and fever. Common gross lesions are characterized by interstitial pneumonia with interlobular septal oedema and major airway oedema. The presence of these gross changes can be seen either in R-PCVAD or S-PCVAD, indicative of an overlap between these two PCV2 presentations (Segalés, 2012). Differentiation between these two clinical presentations is based on histological findings. Characteristic lesions are histiocytic or granulomatous bronchointerstitial pneumonia, peribronchiolar fibroplasia and necrotizing bronchiolitis (Kim et al., 2003; Opriessnig et al., 2007) without the pathognomonic lymphoid lesions observed in S-PCVAD. PCV2 can be detected in interstitial macrophages and bronchiolar lamina propria by IHC and HIS. PCV2 has also been linked with a more severe form of proliferative necrotizing pneumonia (PNP)

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in association with PRRSV, PRV and less frequently SIV (Drolet et al., 2003; Grau-Roma and Segalés, 2007). Histological lesions observed in PNP are characterized by alveolar necrosis and marked hypertrophy and proliferation of type 2 pneumocytes associated with different degrees of lymphohistiocytic interstitial inflammation (Drolet et al., 2003). Although numerous reports suggest the possibility of an overlap between S-PCVAD and R-PCVAD (Harms et al., 2002; Segalés et al., 2005; Opriessnig et al., 2007), recent studies strongly indicate these are not two separate entities, and PCV2 mainly contributes to PRDC in relation to PCV2-SD occurrence (Ticó et al., 2013). Enteric porcine circovirus associated disease (E-PCVAD) Pigs 16–22 weeks old can present diarrhoea caused by PCV2 infection of the intestinal tract. Clinical signs are non-specific and cannot be differentiated from those od other enteric diseases affecting growers and finisher pigs. Diarrhoea is yellow and can progress to dark-red, and is associated with growth retardation and wasting. Morbidity varies from 10% to 20% and mortality is 50–60% (Chae, 2005). The case definition of E-PCVAD is still controversial; however, gastrointestinal lesions have to be associated with the presence of PCV2 in the intestine alone with no other S-PCVAD characteristic or PCV2 detection in other tissues (Segalés, 2012; Baró et al., 2015). The diagnosis of E-PCVAD is appropriate when (Darwich et al., 2004) there is clinical diarrhoea (Vincent et al., 2003). The hallmark microscopic lesions are present in Peyer’s patches but not in other lymphoid tissues (Kekarainen et al., 2010). Antigen or PCV2 DNA or can be demonstrated within lesions (Chae, 2005). Gross lesions are characterized by enlarged mesenteric lymph nodes and thickness of the intestinal mucosa. Histological lesions in E-PCVAD are granulomatous inflammation of the Peyer’s patches characterized by lymphoid depletion and replacement of follicle architecture by macrophages, histiocytes and occasional multinucleated giant cells. The lamina propria and submucosa can also be diffusely infiltrated by macrophages and multinucleated giant cells (Zlotowski et al., 2008). Large, multiple, basophilic or amphiphilic, grape-like intracytoplasmic inclusion bodies are often seen in the cytoplasm of histiocytes and multinucleated giant cells (Kim et al., 2004; Chae, 2005). PCV2 antigen can be detected in histiocytes on Peyer’s patches, submucosa and crypt epithelium by IHC and HIS ( Jensen et al., 2006; Opriessnig et al., 2007). Numerous infections have been observed concomitant with the presence of PCV2. The histological and clinical features of the PCV2 and Lawsonia intracellularis coinfection between have been widely studied (Segalés et al., 2001; Jensen et al., 2006; Opriessnig et al., 2011) in addition to clinical and experimental infections with Salmonella typhimurium (Kim et al., 2004; Opriessnig et al., 2011). Reproductive failure porcine circovirus-associated disease (RF-PCVAD) Reproductive failure associated with PCV2 was first reported in Canada in 1999 (West et al., 1999). Since the initial description, naturally occurring reproductive failures associated with PCV2 have been widely reported (Bogdan et al., 2001; O’Connor et al., 2001; Farnham et al., 2003; Kim et al., 2004; Brunborg et al., 2007). Moreover, experimental studies have also linked PCV2 to reproductive failures ( Johnson et al., 2002; Mateusen, et al., 2004; Pensaert et al., 2004). Although the first reports of S-PCVAD from Canada are dated back from 1991,

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retrospective studies failed to demonstrate the presence of PCV2 in fetal tissues in cases of abortion from the period 1955–1998 (Bogdan et al., 2001). Clinical signs of RF-PCVAD are abortions associated with increased rates of mummified, macerated, stillborn and weak-born piglets (West et al., 1999; O’Connor et al., 2001). However, these clinical presentations can also be the result of numerous infectious and non-infectious causes. Infectious agents including PRRSV, porcine parvovirus (PPV), porcine pseudorabies virus (PRV), and porcine enterovirus can cause almost identical clinical presentations. Non-infectious causes such as mycotoxins, environmental changes or nutritional imbalances might also result in reproductive failures. Coinfection with other pathogens might occur, and numerous pathogens such as PPV (O’Connor et al., 2001; Ritzmann et al., 2005; Pescador et al., 2007), PRRSV (Farnham et al., 2003; Ritzmann et al., 2005), PCV1 (Pescador et al., 2007), have also been reported in association with PCV2; however the significance of these findings is still unknown. PCV2 does not affect specific gestational periods but virus antigen has been detected in early gestation. Therefore PCV2 is associated with embryonic death, irregular returns to oestrus (Kim et al., 2004; Mateusen et al., 2004) in early gestation, with mummified fetuses and abortions (Kim et al., 2004) in mid-gestation, and with mummies, stillborn, weak-born piglets, and delay in farrowing in late gestation (Ladekjaer-Mikkelsen et al., 2001; O’Connor et al., 2001; Johnson et al., 2002). Histological lesions are normally observed in the fetal heart, characterized by lymphocytic infiltration and occasional fibrosis. PCV2 antigen can be detected in the myocardium but can also be observed in other tissues such as liver, kidney and lung (West et al., 1999; Bogdan et al., 2001). Numerous attempts to demonstrate a relation between different PCV2 strain and reproductive failures are documented. Experimental infection of fetuses by intrauterine inoculation (Meehan et al., 2001) or artificial insemination with PCV2 contaminated semen (Madson et al., 2009; Madson et al., 2009) isolated from cases of RF-PCVAD, S-PCVAD, and PDNS, showed no differences in virus replication, tissue tropism and clinical outcomes. Boars can be infected with PCV2 with subsequent viral seeding in semen, making semen an important source of viral dissemination (Larochelle et al., 2000; Kim et al., 2001, 2003). Host immunity and immunopathogenesis Innate immune response The innate, non-adaptive immune response is the perhaps the first barrier against infections as they are activated immediately after infection with a new pathogen. If the pathogen infection is not cleared after the early response, the adaptive immune response takes place. During the early infection, PCV2 has a close interaction with monocytes, macrophages and dendritic cells (DCs) (Darwich et al., 2004). Probably the immune modulation observed during the course of the disease is the result of an early interaction of PCV2 with antigenpresenting cells (APCs). It has been demonstrated that PCV2 can persist in DCs without evidence of virus replication, loss of infectivity or changes in cell viability. PCV2 antigen can persist for several days post infection (dpi) in DCs leading to the conclusion that PVC2 uses DCs infection as a mechanism for viral spread and transmission (Vincent et al., 2003), and the presence of viral particles in those cells is the result of phagocytic or endocytic activity (Steiner et al., 2008; Kekarainen et al., 2010). Clear evidence that other lymphocytes also carry PCV2 is lacking. Any detected PCV2 antigen in lymphocytes could be transient and

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an effect of aberrant cross-talk between the DC carrying the PCV2 and the lymphocyte (Carrasco et al., 2004). No impairment of humoral and cytotoxic T cell responses (Krakowka et al., 2002; Nielsen et al., 2003; Steiner et al., 2009) has been observed in clinically affected pigs, leading to the conclusion that infection of DCs does not affect the immune regulation of lymphocytes by the DCs. Internalization without infectivity has also been observed between PCV2 pulmonary alveolar macrophages (PAMs) and PCV2 monocyte-derived macrophages (MdM) (Gilpin et al., 2003). However, the production of different cytokines in these cells is affected by PCV2 infection. The expression of IL-8 and TNF-α in PAMs, is up-regulated after PCV2 infection (Chang et al., 2006) in association with the increase of mRNA expression of macrophage-derived neutrophil chemotactic factors-II (AMCF-II), granulocyte colonystimulating factor (G-CSF), and monocyte chemotactic protein-1 (MCP-1) (Chang et al., 2006, 2008). In splenic lymphocytes, PCV2 infection decreases IL-4 and IL-2 levels (Darwich et al., 2003a) and increases IL-12 (Duan et al., 2014). There is also down-regulation of IFN-γ, IL-2, IL-4 and IL-12 expression in secondary lymphoid tissue (Darwich et al., 2003b). PCV2 infection of PBMC is modulates proinflammatory cytokines. In PMBCs isolated from PCV2-infected animals, IFN-γ and IL-2 production is impaired (Kekarainen et al., 2008). Up-regulation of proinflammatory cytokines IL-1β and IL-8 has been observed in PBMC from PCV2-infected pigs (Darwich et al., 2003a). In summary, PCV2 can interact with APCs and modulate the immune response without active cellular replication. In addition, PCV2 infection modulates cytokine profiles, which is an important part of the pathogenesis and development of clinical disease. The increase of IL-10 is a common finding in tissues and PBMC of systemic PCVAD affected animals (Darwich et al., 2003a,b; Kekarainen et al., 2008; Crisci et al., 2010; Doster et al., 2010). Overexpression of IL-10 mRNA in multiple lymphoid tissues and cytokine production are important in T-cell rich areas and occasionally in B-cell and macrophages (Doster et al., 2010). Interestingly, this IL-10 stimulation cannot be achieved with Cap or Rep proteins but rather the whole virus is necessary (Crisci et al., 2010). In vivo studies have demonstrated that transient increment of IL-10 in serum is correlated with the viraemic phase observed in subclinically infected animals (Darwich et al., 2008). Adaptive immune response Cell-mediated immune response Studies of cell-mediated immune responses in animals subclinically affected with PCV2 are scarce. IFN-γ-secreting cells (IFN-γ-SC) are the major focus of study in the role of PCV2 and cellular immune response in subclinically infected animals. Studies in caesarean-born and colostrum-deprived pigs (CD-CD) showed that after peak viraemia, coincident with the decreasing serum viral load, there is an increase of PCV2-specific IFN-γ-SC (Fort et al., 2009). These findings support the notion that viral clearance is also due to the contribution of specific IFN-γ-SC in addition to virus neutralization (Fort et al., 2007). In addition, CD4+ and CD8+ lymphocytes seem to play a role in the presence of specific IFN-γ-SC. Depletion of these T-lymphocytes subsets impairs specific IFN-γ-SC (Steiner et al., 2009). Lymphocytic depletion and histiocytic infiltration are the main cellular response observed in clinically affected animals.

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The role of the cell-mediated immune response during clinical presentation is still unclear. However, the lymphocyte impairment observed during clinical presentation suggests that depletion of lymphocytes is a major mechanism of PCV2 pathogenesis. High PCV2 viral loads lead to impairment of neutralizing and T-cell responses (Krakowka et al., 2002; Nielsen et al., 2003; Meerts et al., 2006). Further, up-regulation of IFN-γ-mRNA is observed in PBMCs in clinically affected pigs, which suggests IFN-γ-SC are very important in viral clearance. Similarly, the induction of PCV2-specific IFN-γ-SC are the main cellular component associated with the protective post-vaccination response (Fort et al., 2010; Fort et al., 2012). Humoral response Both the PCV2 capsid and replicase proteins generate antibody responses in infected animals. While the kinetics of the antibody responses to ORF1 have not been studied in detail, cross-reactivity between the ORF1s of PCV1 and 2 is demonstrated and not surprising due to the highly conserved nature of the replicase proteins in circoviruses. Antibodies against the ORF1 do not appear to play an important role in protection, as subunit vaccines containing the capsid protein alone are highly effective against PCV2. However, a strong correlation between ORF2-specific antibodies and protection against PCV2 is well established (Kekarainen et al., 2010). Antibody responses against the PCV2 capsid protein are detected within seven to ten days in infected pigs. However, neutralizing antibody responses are not detected until 14–28 days post infection (Pogranichnyy et al., 2000). In some infected pigs, clinical PCVAD occurs despite high levels of circulating antibodies (Fort et al., 2007). Not all the monoclonal antibodies generated against the PCV2 capsid protein have neutralizing activity, indicating that both conformational, neutralizing and non-neutralizing epitopes are present on the capsid protein (Lekcharoensuk et al., 2004; Constans et al., 2015). Four major immunogenic regions and several linear and conformational epitopes have been identified in the PCV2 capsid protein (Constans et al., 2015). The availability of the PCV2 capsid proteins’ crystal structure has facilitated the prediction of surface residues involved in the interaction with antibodies (Khayat et al., 2011). Further explanation for the delayed protective antibody response is provided by the recent identification of a linear decoy epitope spanning residues 169–180 (Trible et al., 2012). However, monoclonal antibodies or epitope antigens which can distinguish between the various PCV2 strains are not commonly available or used, in the context of protection or serological detection. Polyclonal antibodies from infected pigs show strong cross-reactivity to all known strains of PCV2, indicating serological cross-reactivity and protection between strains. Therefore, the exact mechanisms involved in the protective antibody-mediated immunity against PCV2 are not yet fully characterized. Strong antibody responses are elicited in both infected and vaccinated animals and play an important role in protection against PCV2 (Afghah et al., 2017). Immunosuppression induced by PCV2 It is known that PCV2 affects lymphoid tissues, causing lymphoid depletion followed by granulomatous inflammation with a subsequent immunosuppression during the course of the clinical disease (Rosell et al., 1999; Segalés and Domingo, 2002; Ramamoorthy and Meng, 2009). These histological changes are normally associated with high amounts of PCV2 nucleic acid in affected lymphoid tissues. Lymphoid depletion affects a wide variety

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of lymphoid cells populations including B cells, NK cells, γδ T cells, CD4+ and CD8+ T lymphocytes, and inter-follicular dendritic cells (DCs) (Darwich et al., 2002; Darwich and Mateu, 2012). Although B and T lymphocytes are the most severely affected cells in lymph nodes; PCV2 nucleic acid and antigen have only been found infrequently in lymphocytes (Chianini et al., 2003), instead resident macrophages and APC’s showed strong intranuclear and intracytoplasmic PCV2 signals (McNeilly et al., 1996). However, studies demonstrate that PCV2 can be internalized without evidence of replication in DCs, macrophages and monocytes. The same studies also showed that PCV2 remains infectious and does not induce cell death (Gilpin et al., 2003; Vincent et al., 2003). Virus replication and cell death were not observed in co-cultured DCs and lymphocytes infected with PCV2. Therefore, it has been postulated that viral infection of APCs and subsequent modulation of their function can play a role in viral immune evasion (Vincent et al., 2003). Immune suppression has also been associated with the presence of lymphopenia in clinically S-PCVAD animals, characterized by depletion of CD8+ and CD4+ CD8+ lymphocyte subsets (Darwich et al., 2002). The effect of the presence of PCV2 in lymphocyte populations is still unknown. Several theories have been proposed such as reduced production of bone marrow, reduced proliferation in secondary lymphoid tissues and induction of apoptosis (Shibahara et al., 2000; Resendes et al., 2004; Opriessnig et al., 2007). Based on in vitro data and supported by a mouse model, PCV2 ORF3 has been proposed to be responsible for lymphoid apoptosis (Liu et al., 2006, 2007). The up-regulation of several key factors in the induction of apoptosis has been linked to PCV2 infection. In vitro experiments in PK-15 cells showed an increase on porcine p53 expression, resulting in apoptosis (Liu et al., 2007). PCV2-ORF3 has also been associated with induction of NF-κβ in PK-15 cells and lymphocytes leading to apoptosis (Choi et al., 2015), and the up-regulation of Fas/Fas ligand activity in a PCV2– PRRSV coinfection model. However, the role PCV2 in apoptosis is still debatable because other studies showed that apoptotic rate is inversely correlated with the amount of PCV2 in lymphoid tissues (Resendes et al., 2004). Pigs which were experimentally infected with ORF3 deficient virus did not show differences in lymphoid histological lesions compared to piglets infected with PCV2 wild-type ( Juhan et al., 2010). An alternative theory is that lymphoid depletion occurs due to detriments in cellular proliferation but not apoptosis (Mandrioli et al., 2004). In addition, PCV2 can interfere with the lymphocyte B growth factor, IL-4, and the T cell and macrophage activator IL-2 (Darwich et al., 2003b) which might interfere with lymphocyte proliferation and interferon activity, while increasing the pro-inflammatory cytokines IL-1 and IL-8 (Vincent et al., 2007). While PCV2 does not replicate in DCs (Vincent et al., 2003); viral infection can affect functionality of plasmacytoid dendritic cells (pDCs) (Vincent et al., 2005). Transcription of IFN-α and activation of MyD88, IRF7, IRF3 does not appear to be defective in PCV2 infection (Chen et al., 2016) but the presence of viral DNA, but not viral replication, is sufficient to impair the induction of TNF-α, IFN-α, IL-12 and IL-6 in pDCs (Vincent et al., 2007). The strong up-regulation of IL-10 in PBMCs and PAMs could be responsible for the Th1 suppression (Doster et al., 2010; Du et al., 2016). The results of a PCV2–PRRSV in vitro coinfection model showed an increase in T-reg activation, probably due to IFN-β production but not IL-10 (Cecere et al., 2012; Richmond et al., 2015). A negative synergistic effect has been observed in the mRNA expression of numerous negative regulatory factors associated with toll-like receptors during PCV2–PRRSV coinfection (Dong et al., 2015).

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The role of PCV2 in immunosuppression in S-PCVAD has been widely demonstrated; however, the presence of PCV2 is necessary but not sufficient to develop S-PCVAD. The disease has not been reproduced experientially by singular PCV2 infection; therefore, it is possible that PCV2 needs a cofactor that helps to develop clinical disease. Not even steroid-induced immunosuppression used to mimic PCV2 infections has been successful in reproducing clinical disease. Field studies demonstrated there is a high correlation with clinical PMWS and the presence of PCV2 with multiple coinfecting agents such Mycoplasma hyopneumoniae, Pasteurella multocida, Lawsonia intracellularis, Salmonella spp. and PRRSV, PPV, SIV PSR PEDV, TTV, and porcine Bocavirus (Blomström et al., 2009, 2016). Experimentally, coinfections with multiple bacteria and viruses have been used to reproduce clinical disease. Probably the most efficient and widely used coinfection model is PCV2–PPV. The importance of this model relies not on immunosuppression, as was originally stated but rather immunomodulation (Krakowka et al., 2001; Gillespie et al., 2009; Opriessnig and Halbur, 2012). Interaction amongst multiple pathogens naturally occurs on pig farms; and amongst those the most devastating clinical presentation is observed during PCV2 and PRRSV infection causing severe respiratory disease with a high mortality rate. Clinical diagnosis Direct detection of the virus The clinical diagnosis of PCV2 is rather complex due to the number of different clinical presentations. In addition, the virus is highly prevalent and ubiquitous in most swine production systems. Therefore, the diagnosis of PCV2 has to be evaluated based on clinical context, supported by laboratory diagnosis. Currently, there are numerous techniques available for PCV2 diagnosis. Most laboratories use direct detection methods including conventional polymerase chain reaction (PCR) assays followed by restriction fragment length polymorphism (RFLP), quantitative PCR (qPCR), in situ hybridization and immunohistochemistry (Rosell et al., 1999; Kim et al., 2009). Conventional PCR followed RFLP is still used for subtype identification. Currently cyber green and TaqMan qPCR are the techniques most commonly used for viral quantification in tissues and fluids. In addition, viral sequencing has been used for molecular epidemiology studies and understanding subtype variation in the field. Traditionally, the Sanger method provided sequencing information; howeve,r this method has been replaced by next-generation sequencing techniques (NGS) as it is less time consuming and provides more accurate information. Perhaps the most important feature of NGS is that it can detect nucleic acid from multiple pathogens allowing detection of coinfections. In situ hybridization (ISH) allows a direct correlation with the presence of PCV2 nucleic acid and lesions. Different hybridization approaches and protocols are currently available and most of them target PCV2-ORF2. Specific nucleotide probes can also help to differentiate amongst different PCV2 subtypes (Khaiseb et al., 2011). Immunohistochemistry (IHC) is normally based on monoclonal antibodies generated against PCV2 ORF2, however no subtype differentiation is possible with this technique. Although, the use of immunofluorescence to detect PCV2 in tissues has been replaced by the use of ISH or IHC, this technique is still a viable diagnostic tool for PCV2 in situ detection.

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Serological diagnosis Indirect detection by enzyme-linked immunosorbent assay (ELISA) for antibody detection is widely used to assess exposure to PCV2 or vaccine mediated immunity. Commercial ELISA kits use two detection methods, indirect and blocking formats. The complete ORF2 expressed in baculovirus, whole virus particles, recombinant ORF2 protein and synthetic peptides are used as capture antigens for the ELISAs. Various serological platforms including microbead-based assays such as Luminex have been adapted. Simultaneous detection of antibodies from multiple pathogens make this technique a desirable tool for PCV2 coinfection diagnosis (Lin et al., 2011). All these techniques are capable of detecting PCV2 antibodies generated during natural infection; however, there are no techniques available that differentiate antibodies induced by PCV2 vaccines (DIVA). Perhaps the most difficult clinical presentation to reach an accurate diagnosis is S-PCVAD. By definition an animal or farm with virus circulation detected by PCR, and evidence of infection by presence of PCV2 antibodies should be considered S-PCVAD. However, S-PCVAD diagnosis should not only be supported by laboratory data but also needs to be corroborated by field observations. In the SI-PCVAD clinical presentation, characteristic lymphoid lesions are minimal to none, with low or no nucleic acid in lymphoid tissues detected by ISH/FISH/IHC. There is not a real consensus regarding the viral load necessary to establish the S-PCVAD diagnostic criteria; however several studies have demonstrated that a range of 105 to 106 genomic copy number/ml of serum, without the presence of clinical signs, is consistent with SI-PCVAD (Olvera et al., 2004). The clinical diagnosis of S-PCVAD needs to fulfil three basic criteria. Clinically, the presence of growth retardation, associated with respiratory distress, enlarged lymph nodes and occasionally jaundice that not respond to antibiotic treatment is indicative of S-PCVAD. Demonstration of typical histological lesions of lymphoid depletion, lymphohistiocytic granulomatous lymphadenitis with occasional presence of multinucleated giant cells and characteristic botryoid inclusion bodies is required. Finally aetiological confirmation of PCV2 by IHC or ISH is necessary (Segalés et al., 2005). Herd diagnosis requires the inclusion of both epidemiological and clinical data. A positive S-PCVAD herd definition has been proposed (www.pcv.eu) to include significant increase in post weaning mortality with the presence of clinical signs characteristic of S-PCVAD. Mortality has to be evaluated within the historical levels of the farm. Commonly a consistent increment of two standard deviations from historical data for at least two consecutive production periods is considered significant. In addition, at least one out of five animals should fulfil all the individual diagnostic criteria (Grau-Roma et al., 2011). Since PCV2 infection cause immunosuppression infected animals are prone to secondary and concomitant infections; therefore coinfection with other potential pathogens should to be ruled out for herd diagnosis (Opriessnig and Halbur, 2012). The diagnosis of the PDNS does not necessarily include aetiological diagnosis but is rather based on the presence of characteristic morphological changes and presence of clinical signs. Thus, morphological diagnoses include the presence of macula that progress to necrotizing and haemorrhagic dermatitis. Skin lesions commonly appear in the hind limbs and progress cranially affecting perineum and ventral abdomen. The presence of enlarged, swollen kidneys with multiple cortical petechiae and also enlarged swollen, and haemorrhagic lymph nodes is another diagnostic feature. Histological lesions are characterized by severe necrotizing vasculitis and fibrinous glomerulonephritis. Animals that recovered

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from the disease may present chronic interstitial and proliferative nephritis (Segalés et al., 2005). Therefore, due to the chronic nature of the PDNS lesions, the detection of PCV2 antigen or nucleic acid in situ is rare. Due to the lack of a specific aetiological test a thorough differential diagnosis is necessary in order to rule out other pathogens that cause similar lesions including classical swine fever (CSF), or African swine fever (ASF), both of which are legally notifiable in most countries. Other most commonly observed diseases that need to be differentiated from PDNS include systemic salmonellosis, Erysipelothrix rhusiopathiae and Actinobacillus suis (Drolet et al., 1999). High levels of urea and creatinine, indicative of renal failure, may be detected in serum. The diagnosis of PCV2-RD should be based on the clinical presentation and detection of PCV2 in fetal tissues. Increase in abortions, stillbirth, and mummification in comparison with the historical values in the farm could indicate PCV2-RD (Madson and Opriessnig, 2011). Submission of a group of fetuses is recommended for laboratory diagnosis. Histological lesions of lymphoplasmacytic and necrotizing myocarditis do not provide a definitive diagnosis but can be indicative of PCV2 infection. To confirm the presence of the virus, ISH or IHC of myocardial tissue is recommended (West et al., 1999; Gillespie et al., 2009). Several studies demonstrate that there is a positive correlation between viral load in tissues and the severity of fetal lesions (Brunborg et al., 2007). Although serological evaluation will not demonstrate intrauterine PCV2 infection, testing sows and gilts will help demonstrate previous exposure. Viral load in the serum of sows and gilts have a positive correlation with the PCV2-RD. Since reproductive failures associated with multiple pathogens is common, it is necessary to rule out other reproductive pathogens including PRRSV, PRV and systemic causes of reproductive failure for a definitive PCV2-RD diagnosis. The diagnosis of PCV2 pulmonary disease (R-PCVAD) needs to be included within the evaluation of the typical PRDC workup. The presence of histiocytic interstitial pneumonia needs to be confirmed by viral direct detection by ISH or IHC. In addition detection of the other involved in PRDC would be necessary (Ticó et al., 2013). For the enteric form of PCVAD, the final diagnosis should be based both the clinical features associated with diarrhoea in grower and finisher pigs, but also the presence of lesions that include granulomatous inflammation and lymphocytic depletion in the payer patches, and detection of PCV2 by IHC/ISH. Further, no other lesions or clinical signs indicative of S-PCVAD should to be present (Kim et al., 2004; Opriessnig et al., 2011; Segalés, 2012; Baró et al., 2015). Prevention and control Vaccination Commercial vaccines against PCV2 are highly effective and available in the US and European markets since 2006. The PCV2 capsid protein is considered to be both necessary and sufficient for protection against clinical PCVAD. A majority of the preparations are subunit vaccines containing the PCV2 capsid protein or inactivated PCV1–2 chimeric whole viral particles (Afghah et al., 2017). Vaccination against PCV2 is estimated to increase the average daily weight gain of finishing pigs by about 22g (Alarcon et al., 2013) and result in a cost saving of about $6.00/pig (Gillespie, 2006). In addition, vaccination against PCV2 has substantial benefits in reducing the disease burden due to coinfecting agents, such as PRRSV and SIV (Chae, 2016). All of the current commercial PCV2 vaccines contain the PCV2a

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capsid protein. However, experimental studies suggest that protection against newer, heterologous PCV2b and PCV2d strains, is comparable to the levels of homologous protection elicited by the vaccines (Ssemadaali et al., 2015). Despite such strong experimental evidence, PCV2 continues to evolve in the field and new strains emerge periodically, indicating that the threshold of protection elicited by current vaccines is sufficient to prevent clinical signs but not circulation of the virus and its evolution by recombination or mutation (Ssemadaali et al., 2015). The currently recommended protocol for PCV2 vaccination consists of one to two immunizations between 3 and 6 weeks of age and 2–3 weeks prior to farrowing in sows. While high levels of maternal antibodies will still likely be present in 3- to 6-week -ld piglets, PCV2 vaccines are effective in the presence of maternal antibodies. While there is clear evidence that neutralizing antibody responses are critical for protection, strong vaccine induced antibody responses are not evident in vaccinated pigs with high maternal antibody titres, until eight to ten weeks after vaccination (Dvorak et al., 2017). Therefore, cell-mediated immunity appears to play a role in vaccine-mediated protection, at least in the early vaccine response (Tassis et al., 2017). Current attempts at improving PCV2 vaccines include the development of orally administered vaccines, updated vaccines which include the capsid protein of the newer viral strains and vaccines with markers which can aid in the differentiation of infected and vaccinated animals (DIVA) (Beach et al., 2010, 2011). Biosecurity Despite the effectiveness of PCV2 vaccines and immunization programmes, biosecurity programmes are still critical in preventing the introduction, dissemination, and breaks of S-PCVAD. Typical husbandry practices used in order to reduce S-PCVAD mortality, include environmental management such as temperature, air flow and humidity control. The use of all-in-all-out systems to avoid the mingling of different pigs batches, reduction of crossfostering and, if possible having a solid pen separation to limit animal contact are effective measures to reduce mortality due to S-PCVAD (Madec et al., 2000). Sorting by age and sex at weaning seems also to have an impact on clinical presentation. Weaning animals after 21 days and sorting by sex reduces the risk of S-PCVAD. New introduction of breeding stock including gilts, sows and boars without screening for PCV2 increases the ricks of PCV2 transmission. The presence of PCV2 in semen does not completely imply transmission and the venereal route for transmission is yet to be demonstrated. Although the genetic bases for resistance to PCV2 in certain genetic lines are not known, some genetic lines or pig families have a greater tolerance to infection as they do not present clear clinical signs (Opriessnig et al., 2006). Therefore, the management of the genetic lines is an important point in the control and prevention of clinical PCVAD. Coinfections with other associated pathogens are one of the main triggers of S-PCVAD. Clinical and experimental data has shown that coinfections of PCV2 with porcine reproductive and respiratory syndrome virus (PRRSV), porcine parvovirus (PPV) and Mycoplasma hyopneumoniae can lead to serious losses associated to PCV2 (Rovira et al., 2002). Therefore, vaccination against PCV2 appears to decrease the severity of the coinfections. However, the vice versa is yet to demonstrated. Therefore, adopting a combination of established biosecurity and immunization practices are critical in curtailing clinical PCVAD in production herds.

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Porcine Epidemic Diarrhoea Virus Changhee Lee*

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Animal Virology Laboratory, School of Life Sciences, Kyungpook National University, Daegu, Republic of Korea. *Correspondence: [email protected] https://doi.org/10.21775/9781910190913.05

Abstract A novel enteric disease of swine recognized in Europe in the early 1970s was initially named ‘epidemic diarrhoea’, and is now called ‘porcine epidemic diarrhoea (PED)’. A new coronavirus referred to as PED virus (PEDV) was determined as the etiologic agent of this disease in the late 1970s. PEDV has since plagued Europe and Asia; however, the most severe outbreaks with the greatest economic impact have occurred in Asian swine-producing countries. PED first emerged in the United States in early 2013, caused unprecedented devastation to the pork production industry, and further spread to Canada and Mexico, as well as to South American countries. Promptly thereafter, massive PED epidemics recurred in South Korea, Japan, and Taiwan. Thus, these recent global emergences and re-emergences of PED require urgent attention, and a deeper and concrete understanding of the molecular biology and pathogenic mechanisms underlying PEDV is required to develop effective vaccines and control strategies. This chapter will emphasize the importance of basic, applied, and translational studies and encourage collaboration among swine producers, researchers, and veterinarians to provide answers that improve our knowledge of PEDV in efforts to prevent and eliminate this economically significant viral disease, as well as to prepare for future epizootics or panzootics of PED. History of PEDV (from epizootic to panzootic) In 1971, British veterinarians reported the occurrence of a previously unrecognized enteric syndrome characterized by acute watery diarrhoea in growing and fattening pigs (Oldham, 1972). Representative clinical symptoms of this new type of enteritis were almost identical to those of porcine transmissible gastroenteritis virus (TGEV) infection, and subsequently, the syndrome was named ‘TGE2’. However, in the latter case, neonatal piglets were not affected. The name of this mystery enteric disease was therefore changed to ‘epidemic diarrhoea (ED)’. ED outbreaks extended to multiple swine-producing countries in mainland Europe. After 5 years, TGE-like ED re-emerged in England and also spread to the European

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continent; however, it appeared to be altered. It differed from ED in that this novel diarrheic syndrome now occurred in pigs of all ages including neonatal and suckling animals. Therefore, the ED in 1976 was classified as ED type 2 in order to differentiate it from the initial ED type 1 condition (Wood, 1977; Chasey and Cartwright, 1978). In 1978, researchers at Ghent University in Belgium achieved a breakthrough in ED aetiology, becoming the first research group to fulfil Koch’s postulates, and described the isolation of a new coronavirus-like agent designated CV777 (isolated in July 1977), as the causative pathogen. Furthermore, they provided evidence that this novel virus was distinct from the two known porcine coronaviruses, TGEV and hemagglutinating encephalomyelitis virus (Pensaert and de Bouck, 1978; de Bouck and Pensaert, 1980). Soon thereafter, ED2 was renamed as ‘porcine epidemic diarrhoea (PED)’ caused by the PED virus (PEDV), and has been referred to by this denomination ever since. A marked decrease in acute PED epizootics was observed in Europe in the 1980s and 1990s, and only sporadic outbreaks have occurred in recent years. In these epizootics, adult pigs usually experienced PEDV-associated diarrhoea, whereas suckling piglets were spared or developed only mild symptoms (Saif et al., 2012). PED was first reported in Asia in 1982, and it has since then posed a huge economic threat to the Asian pork industry (Takahashi et al., 1983; Kweon et al., 1993; Chen et al., 2008; Puranaveja et al., 2009; Li et al., 2012). Unlike Europe, PED epizootics in Asia are more severe, causing high mortality in neonatal piglets, and the disease has frequently converted to become enzootic. However, despite its notorious reputation in Asian pork-producing countries, PED was not a globally renowned disease until it hit the United States (US). In May 2013, PED outbreaks suddenly appeared in the USA and spread across the nation rapidly, as well as to adjacent countries. This outbreak caused the death of more than 8 million newborn piglets in the USA alone during a 1-year epidemic period, leading to annual losses in the range of $900 million to $1.8 billion (Mole, 2013; Stevenson et al., 2013; Vlasova et al., 2014; Ojkic et al., 2015; Langel et al., 2016). The USA emergent strain-like viruses further reached East Asian countries, causing nationwide PED disasters (Lee and Lee, 2014; Lin et al., 2014; Lee, 2015; MAFF, 2017). PED has now emerged or re-emerged as one of the deadliest and most contagious viral diseases of swine worldwide, and it is a significant financial menace to the global swine industry. This chapter provides a brief review focusing on current knowledge of the molecular and cellular biology, pathogenesis, diagnosis, and epidemiology of PEDV, as well as control measures to prevent PEDV infection. Molecular biology of PEDV PEDV genome and virion structures PEDV, a large, enveloped, single-stranded, positive-sense RNA virus, is a member of the genus Alphacoronavirus within the family Coronaviridae, placed with three other distantly related families, Arteriviridae, Roniviridae, and Mesoniviridae, in the order Nidovirales based on the similarities in genome organization, predicted proteomes, and replication strategy (Pensaert and de Bouck, 1978; Cavanagh, 1997; Gorbalenya et al., 2006; Saif et al., 2012). The PEDV genome is approximately 28 kb in length with a 5′ cap and a 3′ polyadenylated tail, and consists of seven canonical coronaviral genes, including open reading frame (ORF) 3, in the following conserved order: 5′ untranslated region (UTR)-ORF1a-ORF1b-S-ORF3-E-M-N-3′

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UTR (Kocherhans et al., 2001). The first two large ORFs, ORF1a and 1b, encompass the 5′-proximal two-thirds of the genome and code for non-structural proteins (nsps). ORF1a translation yields a replicase polyprotein (pp) la, whereas ORF1b is expressed by a −1 ribosomal frame shift (RFS) that C-terminally extends ppla into pp1ab. These ppla and pplab are proteolytically matured by internal viral proteases to generate 16 processing end products, named nsp1–16. The remaining ORFs in the 3′-proximal region of the genome encode four canonical structural proteins and one accessory gene, ORF3, individually expressed from the respective 3′-co-terminal nested set of subgenomic (sg) mRNAs, each of which is transcribed from a corresponding sg negative-strand RNA generated by discontinuous transcription of full-length positive-strand genomic RNA. These four structural proteins include the 150–220 kDa glycosylated spike (S) protein, the 20–30 kDa membrane (M) protein, the 7 kDa envelope (E) protein, the 58 kDa nucleocapsid (N) protein, and one accessory gene, ORF3 (Fig. 5.1A) (Duarte et al., 1994; Kocherhans et al., 2001; Lai et al., 2007; Saif et al., 2012; Lee, 2015). The PEDV genome is encapsulated by a single N protein, which forms a long and helical coil structure wrapped in a lipid envelope containing three surface-associated structural proteins, S, M, and E (Fig. 5.1B). Enveloped virions are roughly spherical and pleomorphic, with a diameter ranging from 95 to 190 nm, including the widely spaced, club-shaped, trimerized S projections measuring 18–23 nm in length (Fig. 5.1C) (Pensaert and de Bouck, 1978; Lee, 2015). PEDV has a buoyant density of 1.18 g/ml in sucrose and is stable at 4–50°C. The virus is sensitive to ether and chloroform and is completely inactivated at pH values beyond a pH range of 4–9 (Hofmann and Wyler, 1989). Therefore, a variety of acidic or alkaline chemical disinfectants could destroy PEDV (Pospischil et al., 2002). PEDV structural proteins Major S glycoprotein The S glycoprotein of coronaviruses can be functionally divided into two subdomains, S1 and S2; the former contains a large external portion of S and is responsible for binding to a host-specific receptor, while the latter comprises the remaining external portion, a transmembrane domain, and a short carboxyl terminal cytosolic/virus-internal endodomain, which appears to be involved in direct fusion between the viral and cellular membranes (Lee, 2015). Similar to other coronavirus S proteins, the PEDV S glycoprotein plays a critical role in infection by interacting with its cellular receptor to mediate viral entry and inducing neutralizing antibodies in its natural host ( Jackwood et al., 2001; Lai et al., 2007; Lee et al., 2010; Lee, 2015). Moreover, it is associated with growth adaptation in vitro and attenuation in vivo. Mutations or insertions/deletions in the S gene have been shown to alter viral pathogenicity and tissue/species tropism (Sato et al., 2011). Thus, the PEDV S glycoprotein is an appropriate target viral gene for developing diagnostic assays and effective vaccines (Gerber et al., 2014; Oh et al., 2014). S genes of most PEDV field strains consist of 4,161 nucleotides (nt) encoding 1386 amino acid (aa) residues, which are 9-nt (3-aa) longer than the homologous gene of the prototype CV777 strain. Compared with the sequence of CV777, PEDV epidemic strains possess distinct genetic signatures, namely S insertions-deletions (S INDELs) comprising two notable 4-aa and 1-aa insertions at positions 55/56 and 135/136, respectively, and a unique 2-aa deletion at positions 160 and 161 (Lee et al., 2010) (Fig. 5.2). Although only

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A

B

Figure 5.1  Schematic representations of PEDV genome organization and virion structure. (A) The structure of PEDV genomic RNA. The 5′-capped and 3′-polyadenylated genome of approximately 28 kb is shown at the top. The viral genome is flanked by UTRs and is polycistronic, harbouring replicase ORFs 1a and 1b followed by the genes encoding the envelope proteins, the N protein, and the accessory ORF3 protein. S, spike; E, envelope; M, membrane; N, nucleocapsid. ORF1a and 1b expression yields two known polyproteins (pp1a and pp1ab) by –1 programmed RFS; these polyproteins are co-translationally or posttranslationally processed into at least 16 distinct non-structural proteins designated nsp1–16 (bottom). PLpro, papain-like cysteine protease; 3CLpro, the main 3C-like cysteine protease; RdRp; RNA-dependent RNA polymerase; Hel, helicase; ExoN, 3′5′ exonuclease; NendoU, nidovirus uridylate-specific endoribonuclease; 2′OMT, ribose-2′-O-methyltransferase. (B) Model of PEDV structure. The structure of the PEDV virion is illustrated on the left. The RNA genome inside the virion is associated with the N protein to form a long, helical ribonucleoprotein (RNP) complex. The virus core is enclosed by a lipoprotein envelope, which contains S, E, and M proteins. The predicted molecular sizes of each structural protein are indicated in parentheses. A set of corresponding subgenomic mRNAs (sg mRNA; 2–6), through which canonical structural proteins or non-structural ORF3 protein are exclusively expressed via a co-terminal discontinuous transcription strategy, are also depicted on the right. Adapted from Lee (2015). (C) Electron micrograph of a PEDV particle showing typical coronavirus morphology. The arrow points to virus peplomers or spikes. Bar = 100 nm.

one PEDV serotype has been reported, phylogenetic studies of the S gene suggested that PEDV can be genetically separated into two genogroup clusters, genogroup 1 (G1, classical and recombinant: low pathogenic) and genogroup 2 (G2, field epizootic or panzootic: high pathogenic), which are further divided into subgroups 1a and 1b, and 2a and 2b (Fig. 5.2). G1a includes the prototype CV777, vaccine, and other cell culture-adapted strains, whereas G1b comprises new variants that were first identified in China (Li et al., 2012), later in the USA (Wang et al., 2014) and South Korea (Lee et al., 2014b), and recently in European countries (Hanke et al., 2015; Theuns et al., 2015; Grasland et al., 2015). G1b strains are assumed to arise from homologous recombination between classical G1a and field G2 viruses. G2 contains global field isolates, which are clustered into 2a and 2b subgroups

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Figure 5.2  Phylogenetic tree based on the spike nucleotide sequence of global PEDV strains and the amino acid sequence alignment of the N-terminal region of the S protein of global PEDV strains. Multiple sequence alignments were performed using ClustalX, and the phylogenetic tree was constructed from aligned nucleotide sequences using the distance-based neighbourjoining method. Numbers at each branch represent bootstrap values greater than 50% of 1000 replicates. Only the corresponding alignment of amino acid sequences of the N-terminal region containing hypervariable regions (Lee et al., 2010) is shown, and amino acid sequences of the prototype CV777 strain are shown at the top. Genetic subgroups of PEDV are indicated by different colours: G1a (red), G1b (blue), G2a (green), and G2b (purple). Asterisks (*) indicate mutated sequences. Insertions and deletions (INDELs) within PEDV isolates compared to the prototype CV777 strain are shaded. Asterisks (*), dashes (-), and pluses (+) indicate mutated, deleted, and inserted sequences, respectively, compared with CV777.

responsible for the past regional epidemics in Asia, and for the 2013–2014 pandemics and current outbreaks in the American and Asian continents, respectively. Considering those genetic traits, the S gene could be a suitable locus for sequencing to investigate genetic relatedness and molecular epidemiology of PEDV (Lee et al., 2010, 2014a,b; Chen et al., 2014; Lee and Lee, 2014).

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Other structural and species-specific (accessory) proteins The M protein, the most abundant component of the viral envelope, is required for the assembly process, and can also elicit production of protective antibodies with virus-neutralizing activity (de Hann et al., 2000; Zhang et al., 2012). The small envelope E protein plays an important role during coronavirus budding, and co-expression of E and M proteins can form spike-less coronavirus-like virions (Baudoux et al., 1998). PEDV E and N proteins are found in the endoplasmic reticulum (ER), independently inducing ER stress (Xu et al., 2013a,b). The N protein has multiple functions in viral replication and pathogenesis in coronavirology (McBride et al., 2014). Generally, coronavirus N proteins interact with viral genomic RNA and associate with other N protein molecules to protect the viral genome, serving as the critical basis for the helical nucleocapsid during coronavirus assembly (McBride et al., 2014). The PEDV N protein also perturbs antiviral responses by antagonizing interferon production as part of the immune evasion strategy, and activates NF-κB signalling (Xu et al., 2013b; Ding et al., 2014). The product of ORF3, the sole PEDV accessory protein, functions as an ion channel that influences virus reproduction and virulence (Wang et al., 2012). Furthermore, a large deletion in the ORF3 region is present in attenuated or live vaccine strains (Park et al., 2008; Wang et al., 2012). Inconsistent data provide evidence that ORF3 disables recovery of the recombinant progeny virus rescued from a full-length infectious clone bearing no ORF3 start codon, suggesting that functional ORF3 negatively modulates PEDV replication in vitro ( Jengarn et al., 2015). However, studies revealed that the PEDV ORF3 product is dispensable for the replication of the virus in cultured cells, indicating its ignored function in virus propagation in vitro (Li et al., 2013; Lee et al., 2017). Nevertheless, conservation of the complete ORF3 in PEDV field isolates suggests that the ORF3 protein plays a critical role in natural infection and pathogenesis in the animal host. Cellular biology of the virus Cell tropism Coronaviruses infects a wide range of mammals, including humans, bats, and whales, as well as birds, but they typically have a limited host range, infecting only their specific natural host. Furthermore, coronaviruses exhibit a marked tropism for epithelial cells of the respiratory and enteric tracts, as well as for macrophages (Woo et al., 2012; McVey et al., 2013; Reguera et al., 2014). Likewise, PEDV shows restricted tissue tropism and replicates efficiently in porcine small intestinal villous epithelial cells or enterocytes. Porcine aminopeptidase N (pAPN), predominantly expressed on the epithelial cell surface of the small intestine, has been identified as the cellular receptor for PEDV (Li et al., 2007; Nam and Lee, 2010). The N-terminal region of the PEDV spike protein S1 domain is important for recognizing the pAPN receptor (Lee et al., 2011). However, most recent studies report that the APN is not a functional cellular receptor for PEDV, suggesting the presence of the authentic virus receptor essential for viral entry (Shirato et al., 2016; Li et al., 2017). Additionally, cell-surface heparan sulfates have been shown to act as the attachment factor for PEDV (Huan et al., 2015). Nevertheless, PEDV entry begins with binding to pAPN or a yet-to-be-identified main receptor, followed by virus internalization into target cells by direct membrane fusion, and subsequent release of the viral genome into the cytosol after uncoating to start genome

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replication (Fig. 5.3). In addition to replicating in the natural host’s primary target cells, PEDV can grow in porcine cells isolated from different organs, or cells derived from species other than swine, including those from monkeys, such as African green monkey kidney cell lines (Vero and MARC-145), ducks, bats, and humans (Hofmann and Wyler, 1988; Lawrence et al., 2014; Khatri, 2015; Liu et al., 2015; Wang et al., 2016). Among these, Vero cells are generally accepted to be the most suitable for PEDV isolation and propagation. The actual cellular receptor for PEDV appears to be evolutionarily conserved among the

Figure 5.3  PEDV replication and pathogenesis. PEDV binds pAPN or a hitherto-unidentified cellular receptor via the spike protein on villous epithelial cells of the small intestine. The S protein-mediated fusion of the viral envelope with the plasma membrane is followed by penetration and uncoating of the virus. Following disassembly, the viral genome is released into the cytoplasm and immediately translated to yield the replicases, ppla and pp1ab. These polyproteins are proteolytically cleaved into 16 nsps comprising the replication and transcription complex (RTC) that first engages in the minus-strand RNA synthesis using genomic RNA. Both full- and sg-length minus strands are produced and used for synthesis of full-length genomic RNA and sg mRNAs. Each sg mRNA is translated to yield only the protein encoded by the 5′-most ORF of the sg mRNA. The envelope S, E, and M proteins are inserted in the ER and anchored in the Golgi apparatus. The N protein interacts with newly synthesized genomic RNA to form helical RNP complexes. The progeny virus is assembled by budding of the preformed RNP at the ER-Golgi intermediate compartment (ERGIC) and then released by the exocytosis– like fusion of smooth-walled, virion-containing vesicles with the plasma membrane (Lai et al., 2007). Adapted from Lee (2015). PEDV infection destroys target enterocytes, resulting in villous atrophy and vacuolation. This consequence disrupts digestion and absorption of nutrients and electrolytes, thereby causing acute maldigestive and malabsorptive watery diarrhoea and ultimately leading to serious and fatal dehydration in neonates.

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aforementioned species, and its identification is critical for the development of control strategies to prevent PEDV. Although whether APN acts as the functional receptor for PEDV on these cells still remains to be determined, overexpression of exogenous pAPN renders non-permissive cells susceptible to PEDV infection. This observation suggests a functional significance of the APN density for PEDV propagation in cell culture (Nam and Lee, 2010). Furthermore, pAPN promotes PEDV infection via its enzymatic activity, indicating an irrelevant role of pAPN in viral entry as a receptor during PEDV replication (Shirato et al., 2016). Supplement with trypsin is indispensable for isolation and serial cultivation of PEDV in Vero cells (Hofmann and Wyler, 1988; Chen et al., 2014; Oka et al., 2014; Lee et al., 2015). Trypsin facilitates PEDV entry and release by cleaving the S protein into S1 and S2 subunits, enabling successful viral replication and spread in vitro (Shirato et al., 2011; Wicht et al., 2014). However, some cell-adapted attenuated PEDV strains, such as SM98-1 and 83P-5, can support PEDV propagation in the absence of trypsin (Nam and Lee, 2010; Kim and Lee, 2013). As a result of viral infection, distinct cytopathic effects (CPEs) including cell fusion, vacuolation, syncytium, and detachment are developed in a stepwise manner in infected cells (Lee et al., 2015). PEDV–host interactions Since viruses rely on the host machinery to make new viruses, they might adjust the activity of host cellular factors or signalling pathways to benefit their own multiplication. Proteome analysis revealed that the expression of proteins involved in apoptosis, signal transduction, and stress responses is modulated in PEDV-infected Vero cells (Zeng et al., 2015). PEDV induces apoptotic cell death in vitro and in vivo through the caspase-independent mitochondrial apoptosis-inducing factor (AIF) pathway that plays a critical role in PEDV replication and pathogenesis (Kim and Lee, 2014). PEDV infection activates the three major mitogenactivated protein kinase (MAPK) cascades involving extracellular signalling-regulated kinase (ERK), p38 MAPK, and c-Jun N-terminal kinase ( JNK) (Kim and Lee, 2014, 2015; Lee et al., 2016). Additionally, PEDV appears to induce ER stress and activate NF-κB in infected cells (Xu et al., 2013a,b). Therefore, viral replication and subsequent pathological changes depend on the ability of PEDV to exploit multiple intracellular processes, such as apoptosis, MAPK signalling, and ER stress, which emerge in response to various extracellular stimuli. Upon viral infection, a host immediately reacts to the invading virus by producing type I interferon (IFN), which is a key mediator of innate antiviral responses. To counteract innate immune signalling, several viruses, including coronaviruses, have evolved different strategies to modulate the activation of antiviral cytokines forming host innate immunity, particularly by diminishing IFN induction and/or inhibiting IFN signalling (Perlman and Netland, 2009). Like other coronaviruses, PEDV has developed certain mechanisms to circumvent the IFN response by limiting IFN production, and encodes several structural and non-structural proteins that serve as type I IFN antagonists (Xing et al., 2013; Cao et al., 2015; Zhang et al., 2016; Zhang and Yoo, 2016). PEDV also antagonizes the antiviral effect of IFN by suppressing type I IFN signalling pathways in response to degradation of the signal transducer and activator of transcription (STAT) protein 1 in a proteasomedependent manner (Guo et al., 2016). Thus, PEDV evades the host innate immunity by subverting the type I IFN synthesis and signalling pathways, which in turn, contributes to viral replication and pathogenesis.

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Viral pathogenesis Transmission The spread of PEDV within populations of pigs and between farms occurs principally by a direct or indirect faecal–oral route; no vector or reservoir has been implicated in its transmission (Fig. 5.4). PEDV can mainly invade farms via diarrhoeal faeces or vomitus, and through contaminated environmental sources via clinically or subclinically infected pigs, trailers transporting pigs, manures, or food, rendering trucks, humans (pig owners or visitors, such as swine practitioners or trailer drivers wearing contaminated work clothing and footwear), wild animals including birds, or insects (Saif et al., 2012; Lowe et al., 2014). Other contaminated fomites such as sow milk, feed, food items, or food additives or ingredients, including spray-dried porcine plasma, could all be potential sources of PEDV transmission (Li et al., 2012; Dee et al., 2014; Opriessnig et al., 2014; Pasick et al., 2014). Airborne transmission may also play a role in PEDV dissemination under certain conditions (Alonso et al., 2014). In general, the virus may vanish following acute PEDV outbreaks, or remain in the farrowing unit because of inadequate hygiene management (e.g. improper disinfection and lax biosecurity), or it may persist in weaning or growing-finishing facilities, causing mild post-weaning diarrhoea without mortality (Lee, 2015). In the latter endemic status, if newborn piglets are unable to obtain ample levels of maternal immunity from their dams due to incomplete sow vaccination or defective lactation owing to mastitis or agalactia, the virus that has circulated in the farm will infect susceptible piglets, serving as the source of recurrence of epidemic outbreaks, and ultimately resulting in an explosive increase in the death of neonatal pigs (Park and Lee, 2009; Park et al., 2011). Although such endemic PED circumstances are frequently reported in Asian countries where PED epidemics occur and recur repeatedly, they may likewise happen in newly PEDV emerging nations, including the USA.

Figure 5.4  PEDV transmission sources and routes in epidemic and endemic cases.

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Clinical and pathological observations PEDV can infect pigs of all ages, causing anorexia and depression accompanied by vomiting and watery diarrhoea. Morbidity approaches 100% in piglets, but can vary in adult pigs (Saif et al., 2012). The PEDV incubation period is approximately 2 days, ranging from 1 to 8 days, depending on field or experimental conditions. The interval between the onset and cessation of clinical signs is three to four weeks (Pensaert and de Bouck, 1978; Saif et al., 2012; Wang et al., 2013; Jung et al., 2014; Madson et al., 2014; Lee et al., 2015). Faecal shedding of PEDV can be detected as early as 24 h post infection and may last for up to 4 weeks. The disease severity and mortality rates might, however, be inversely associated with the age of the pigs (Shibata et al., 2000; Saif et al., 2012). PEDV infection in neonatal piglets up to 1 week of age causes severe scours and vomiting for 3–4 days, followed by extensive dehydration and electrolyte imbalance, eventually leading to death (Fig. 5.5). The mortality rate averages 50%, often approaching 100% in 1- to 3-day-old newborn piglets, and decreases gradually to 10% thereafter. In older animals, including weaner to finisher pigs, clinical symptoms are self-limiting within the first week after the onset of the disease. However, PED may affect growth performance of growing pigs. Sows may not experience diarrhoea, but often manifest depressive symptoms and anorexia. If farrowing sows lose their offspring, they may subsequently suffer from reproductive disorders including agalactia, delayed estrus, or reduced pregnancy rate and litter size, which result from the absence of suckling piglets during the lactation period. Gross lesions are confined to the gastrointestinal tract and are characterized by a distended stomach filled with completely undigested milk curd and thin, transparent intestine walls with the accumulation of yellowish fluids (Fig. 5.6A) (de Bouck et al., 1981; Jung et al., 2014; Lee et al., 2015). Histological hallmarks of the PEDV infection include severe diffuse

Figure 5.5  Representative clinical symptoms of PEDV infection. (A) Piglet watery diarrhoea or scours (arrows). (B) Runt piglets. (C) Mass mortality. (D) Mastitis in a lactating sow.

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Figure 5.6  Representative macroscopic and microscopic lesions of the intestine from piglets infected with PEDV. (A) Small intestines with thin and transparent intestinal walls. (B) Haematoxylin and eosin-stained jejunum with villous vacuolation. (C) Haematoxylin and eosinstained jejunum exhibiting villous atrophy and exfoliation.

atrophic enteritis, superficial villous enterocyte swelling with mild cytoplasmic vacuolation, necrosis of scattered enterocytes followed by sloughing, and contraction of the subjacent villous lamina propria containing apoptotic cells (Fig. 5.6B) (de Bouck et al., 1981; Jung et al., 2014; Madson et al., 2014; Lee et al., 2015). The intestinal villi become shortened to two-thirds or less of their original length (villous height to crypt depth ratios change to less than 3 : 1 in affected pigs) with the extent of the pathology depending on the infection or disease process stage (Fig. 5.6B) (de Bouck et al., 1981; Jung et al., 2014; Madson et al., 2014). Pathogenesis PEDV replicates in the cytoplasm of villous epithelial cells throughout the small intestine, destroying target enterocytes, possibly because of massive necrosis or apoptosis (Kim and Lee, 2014). Consequently, PEDV infection causes villous atrophy and vacuolation, as well as a significant reduction in enzymatic activity (Saif et al., 2012; Kim and Lee, 2014). The sequence of the disease process hinders digestion and absorption of nutrients and electrolytes, thereby resulting in malabsorptive watery diarrhoea, followed by serious and fatal dehydration in neonatal piglets (Ducatelle et al., 1982; Saif et al., 2012). Upon infection with PEDV, the disease outcome and PED-associated deaths usually occur in an age-dependent manner. Although the reasons for induction of more severe disease in nursing piglets in comparison to weaned pigs by PEDV have not been definitively elucidated, slower regeneration of enterocytes in neonatal pigs may be an important factor (Moon et al., 1973). PEDV infection increases the number of crypt stem cells and the proliferation of crypt cells, pointing to accelerated epithelial cell renewal ( Jung and Saif, 2015). Enterocyte turnover rate was slower in normal nursing piglets than in weaned pigs, suggesting that the speed of crypt stem cell replacement is associated with age-dependent resistance to PED ( Jung and Saif, 2015). Diagnostic procedures Because the outcomes of PEDV infection are clinically and pathologically indistinguishable from those caused by other porcine enteric coronaviruses including TGEV and porcine deltacoronavirus (Haelterman, 1972; Ducatelle et al., 1982; Ma et al., 2015), PED diagnosis

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cannot be made solely based on clinical symptoms and pathological lesions. Therefore, differential diagnosis to detect the presence of PEDV at its antigen or/and genome levels must be conducted in the laboratory. Various traditional virological methods, including immunofluorescence (IF) or immunohistochemistry (IHC) tests, in situ hybridization, electron microscopy, virus isolation, enzyme-linked immunosorbent assays (ELISA), and reverse-transcription polymerase chain reaction (RT-PCR) techniques, have been applied (Fig. 5.7). Taking their rapid turnaround times and sensitivity into account, conventional and real-time RT-PCR systems available as commercial kits are most widely used for PEDV diagnosis during epidemic or endemic outbreaks, as well as for quarantine or slaughter policies. In addition, nucleotide sequencing of the S gene region may be useful for determining the PEDV genotype circulating in herds. The combination of RT-PCR and S gene sequencing could prove to become an optimal tool for diagnosing the virus and surveilling the emergence of novel variants in the field (Lee, 2015). Numerous serological assays have been utilized for the detection of PEDV antibodies, including indirect fluorescent antibody (IFA) staining, ELISA, and virus neutralization

Figure 5.7  Laboratory methods for diagnosis of PEDV infection. (A and B) IHC or IF detection of PEDV antigens in jejunum. Brown (A) or green (B) indicates PEDV N antigen in the cytoplasm of infected enterocytes. Antigens were detected with anti-PEDV N protein monoclonal antibody (MAb) (CAVAC, Daejeon, South Korea). (C) Conventional RT-PCR for detection of PEDV nucleic acid in clinical samples such as faeces, rectal swabs, or small intestinal homogenates. The PEDV genome was detected by RT-PCR assay using an i-TGE/PED Detection Kit (iNtRON Biotechnology, Seongnam, South Korea). The last lane represents a positive control for TGEV (upper band) and PEDV (lower band). (D–F) Isolation of PEDV in cell culture. PEDV-specific CPEs characterized by syncytia and cell detachment (D), multinucleated cells (polykaryon) stained with DAPI (E), and IFA confirming virus isolation using a PEDV N-specific MAb (F) in Vero cells.

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(VN) tests. Due to the special protection strategy (passive immunization) for neonatal piglets against PEDV, determining the presence or absence of anti-PEDV antibodies may be meaningless in sow herds. Instead, measuring quantities (or titres) of neutralizing antibodies (NA) against PEDV or, especially, against the S protein in colostrum and sow milk and/or serum, should be necessary for monitoring immunity levels following sow immunization. In this regard, the VN test could be essential for estimating levels of protective antibodies that newborn piglets would receive from the sows. However, this method is time-consuming, and cannot selectively detect only secretory IgA antibodies representing mucosal immunity (Lee, 2015). In contrast, IFA and indirect ELISA approaches for antibody detection are equally specific but less time-consuming and easier to perform compared to the VN test. Most assays that are currently in use have been developed based on either the whole virus (Hofmann and Wyler, 1990; Carvajal et al., 1995a; Oh et al., 2005) or viral protein antigens (Knuchel et al., 1992; Gerber et al., 2014). Whole-virus-based IFA and ELISA tests may be inappropriate for evaluating the protective NA because they also capture various antibodies against virion components such as M or N proteins. However, these tools may still be useful for monitoring PEDV enzootic situations in affected farms by determining the infection status in weaner to finisher pigs. On the other hand, the entire S protein or its S1 portion could be suitable for viral antigens used in ELISA because the S1 domain of PEDV contains the potential receptor-binding region and main neutralizing epitopes (Sun et al., 2007; Lee et al., 2011). Indeed, a recombinant S1 protein-based indirect ELISA has been developed to detect anti-PEDV antibodies (Gerber et al., 2014). Furthermore, Song et al. (2016) showed that the neutralizing activity against PEDV in mammary secretions significantly correlated with IgA primarily specific to S1 and S2 during lactation. Thus, VN and S1-based ELISA methods are recommended for assessing the protective capacity of sows in their colostrum and milk, which are critical for suckling neonates to acquire early passive protection against PEDV. Molecular epidemiology Epidemiology in Europe Despite its first appearance in the United Kingdom, and later in multiple European countries in the 1970s, the impact and financial importance of PEDV in Europe are currently negligible compared with those in Asian countries and in the USA. Therefore, the epidemiology of PEDV in Europe over the past decades has not been studied intensively. In the 1980s and 1990s, PEDV outbreaks became infrequent, while the virus persisted endemically in the pig population at a low prevalence rate. Sporadic outbreaks were reported in some European nations, causing diarrhoea in weaner or feeder pigs. Numerous serological surveys indicated that seroprevalence of PEDV became low in European pigs (Pijpers et al., 1993; Van Reeth and Pensaert, 1994; Carvajal et al., 1995b; Nagy et al., 1996; Pensaert and Van Reeth, 1998; Pritchard et al., 1999). Interestingly, even although herd immunity against PEDV in European countries is deficient, the virus rarely caused severe outbreaks in these susceptible pig populations, and, accordingly, the exact resistance mechanism remains to be elucidated. In 2006, a typical epidemic form of PEDV re-emerged among pigs of all ages in Italy (Martelli et al., 2008). A case of PED on a fattening farm was reported in Germany in 2014 (Hanket et al., 2015). Shortly thereafter, outbreaks of PEDV were identified in a farrow-finish herd

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in France and in fattening pigs in Belgium (Theuns et al., 2015; Grasland et al., 2015). These German, French, and Belgian PEDV strains were found to be nearly genetically homologous to each other, and most closely resemble the G1b variants discovered in China, the USA, and South Korea (Fig. 5.3). Further surveillance studies are needed to determine whether the G1b strains were previously circulating in Europe, or if they were recently introduced from the USA or Asia. Around the same time, a PEDV epidemic occurred in Ukraine, severely affecting nursing piglets with high fatality rates, and pandemic G2b-like strains were found to be responsible for this emergence (Dastjerdi et al., 2015). This PED outbreak highlights the threat to neighbouring and more distant nations in the European Union. Therefore, the implementation of strict biosecurity protocols would be necessary to prevent the further spread of PEDV domestically or internationally in Europe. In particular, it would be important to continue monitoring the situation if the highly virulent G2 PEDV is present in certain areas of Europe, where the virus has not emerged or re-emerged in recent decades. Epidemiology in Asia In Asia, PED epidemics first occurred in 1982 in Japan and, since then, PED has caused severe epidemics in adjacent Asian countries, particularly in China and South Korea, resulting in heavy losses of piglets (Takahashi et al., 1983; Kweon et al., 1993; Jinghui and Yijing, 2005). PED has remained rampant, leading to serious economic losses in China since its first identification. In the early 1990s, a vaccine containing the inactivated prototype CV777 strain was developed, and has since been widely used throughout the swine industry in China. Until 2010, PED outbreaks were intermittent with only a limited number of incidents. However, a remarkable increase in PED epidemics occurred in pig-producing provinces in late 2010 (Li et al., 2012). During this period, new PEDV variants belonging to the G1b genogroup were first reported in China (Li et al., 2012). Since then, intense PEDV epidemics have been reported in various regions of China, and both G1b variants and field epidemic G2 strains have been responsible for current PED outbreaks in China (Sun et al., 2012; Wang et al., 2013; Tian et al., 2014). A G2b strain, AH2012, was later found to be a potential progenitor of USA PEDV emergent strains in 2013 (Huang et al., 2013; Vlasova et al., 2014). In the late 2000s, PEDV was reported and became increasingly problematic in the Philippines, Thailand, Taiwan, and Vietnam (Puranaveja et al., 2009; Duy et al., 2011; Lin et al., 2014). PEDV isolates responsible for epidemics in Thailand had S genetic signatures typical for field epidemic G2 strains, and were placed in the cluster adjoining South Korean and Chinese strains in the G2a or G2b subgroup (Temeeyasen et al., 2014). PED was first observed in southern provinces of Vietnam and soon after, the disease spread throughout all major swine-producing regions (Duy et al., 2011). Vietnamese strains also had unique S INDEL characteristics and could be classified as the G2b sublineage, which continues to cause sporadic outbreaks in Vietnam (Vui et al., 2014). Prior to late 2013, PEDV incidence was relatively low with only sporadic outbreaks in Taiwan and Japan. In late 2013, severe large-scale PED epizootics suddenly re-emerged in these countries, which led to tremendous financial losses in their pork industry (Lin et al., 2014; MAFF, 2017). Taiwanese and Japanese viral isolates collected in 2013 to 2014 were phylogenetically related to the same clade as global G2b PEDV strains (Lin et al., 2014; MAFF, 2017).

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Epidemiology in the United States PEDV was exotic in the USA until its unexpected and explosive emergence in May 2013. Since then, PEDV has spread swiftly in swine farms across the USA, posing significant financial threats (Stevenson et al., 2013). Genetic and phylogenetic analyses of the emergent US PEDV strains identified during the initial outbreak revealed a close relationship with Chinese strains, especially the AH2012 strain isolated in 2012 from Anhui Province in China, implying their origin (Huang et al., 2013). Recent studies suggested that the emergent PEDV strains in the USA potentially descended from two Chinese strains, AH2012 and CH/ZMDZY/11, in the G2b sublineage, through recombination (Vlasova et al., 2014; Tian et al., 2014). Remarkably, PEDV strains similar to those found in the USA appear to be responsible for subsequent large-scale PED outbreaks in South Korea, Taiwan, and Japan in late 2013 (Lee and Lee, 2014; Lin et al., 2014; MAFF, 2017). Subsequently, other novel US PEDV strains, such as OH851, without S protein genetic signatures typical of the epidemic G2 virus, were reported in 2014. They were phylogenetically clustered closely to novel Chinese strains in the G1b subgroup based on similarities of the S gene, or with the emergent US PEDV strains in the G2 group based on whole genome characteristics (Wang et al., 2014). Novel variants from the USA had a low nucleotide identity in the first 1,170 nucleotides of their S1 region, and a high similarity in the remainder of the S gene, compared to the PEDV strains mainly circulating in the USA, suggesting a rapid evolution of US PEDV variants through potential recombination events (Wang et al., 2014). However, a retrospective study demonstrated that the new US variants were already present in June 2013, indicating a possibility that multiple parental PEDV strains were introduced into the USA at approximately the same time (Vlasova et al., 2014). Epidemiology in South Korea The first PED epizootic in South Korea was confirmed in 1992 (Kweon et al., 1993). However, a retrospective study revealed that PEDV already existed in South Korea since as early as 1987 (Park and Lee, 1997). Since then, PED outbreaks have occurred annually and have become endemic, which resulted in high rates of piglet death and substantial economic losses to the domestic swine industry until 2010. In a serological survey conducted in 2007, 91.8% of 159 tested farms had sero-positive pigs in wean-to-finish periods (30–150 days of age), indicating that the majority of farms were heavily affected by endemic PEDV infection (Park and Pak, 2009). Since the early 2000s, both modified live and inactivated vaccines, based on domestic isolates SM98-1 or DR-13, have been introduced nationwide, leading to a decline in the incidence of PEDV-associated diarrhoeal disease outbreaks compared to previous years (Lee, 2015). However, continuous PED epidemics in vaccinated farm animals have raised questions about the efficacy of Korean commercial vaccines. The dominant PEDV isolates in South Korea during the same period were classified as G2a strains with S INDEL signatures, and were distantly related to prototype CV777 or Korean vaccine strains belonging to the G1a subgroup (Lee et al., 2010). Since South Korea underwent severe outbreaks of foot-and-mouth disease (FMD) in 2010–2011, there was a state of lull in PED epidemics. The prevalence of PEDV infections was occasional, with only intermittent outbreaks in South Korea from 2011 to early 2013. This epidemic situation likely resulted from the mass culling of more than 3 million pigs (onethird of the entire domestic pig population) in South Korea during the 2010–2011 FMD outbreaks (Lee, 2015). However, from November 2013 onwards, severe PED epidemics

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increased dramatically and swept through nearly half of the pig farms across mainland South Korea (Lee and Lee, 2014); 4 months later, the virus arrived in Jeju Island, which had been free of PEDV since 2004 (Lee et al., 2014a). The re-emergent PEDV isolates responsible for massive epidemics in South Korea in 2013–2014 were classified into the G2b subgroup, and were clustered closely with emergent USA PEDV strains (Lee and Lee, 2014; Lee et al., 2014a). The source of PEDV incursion into the South Korean swine population has not yet been determined. The import of pig breeding stock during or after the sudden emergence of PEDV in the USA might be a possible sources, but it remains unclear whether G2b PEDVs similar to USA strains were pre-existing in South Korea. Indeed, two Korean G2b isolates, KDJN12YG and KNU-1303, were identified independently in November 2012 (Kim et al., 2015) and May 2013 (Lee and Lee, 2014), respectively. The former was similar to Chinese G2b strains, while the latter was similar to emergent USA G2b strains. Hence, it is also conceivable that the virus, which has evolved independently by recombination or point mutations, might have already been present in South Korea as a minor lineage before the PED emergence in the USA (Lee, 2015). Alternatively, it might have originated directly from China, and suitable circumstances have since made G2b strains dominant, leading to a number of recent acute outbreaks nationwide (Lee and Lee, 2014; Kim et al., 2015; Lee, 2015). Novel variant G1b PEDV isolates were found in South Korea in March 2014, and these were analogous to the variants reported in China, the USA, and recently, in several European countries (Lee et al., 2014b). They had common genetic and phylogenetic features of G1b strains (no S INDELs compared to CV777, different phylogenetic subgroup [G1b or G2] depending on the sequence of the S protein or whole-genome, and evidence of recombination), and, among other G1b strains, were most closely related to the USA variant strain OH851 among other G1b strains (Lee et al., 2014b). Although a temporal study will be needed to verify the presence of the G1b virus in earlier periods prior to its first identification, it is feasible that, as in the case of outbreaks in the USA, two G1b and G2b ancestor strains resembling USA strains could have been simultaneously transmitted into South Korea (Lee, 2015). Another novel PEDV G2 strain (MF3809) with a large S deletion was found in South Korea. However, this isolate was identified from only three diarrhoea samples out of 2634 on one out of the 569 farms surveyed in 2008 (Park et al., 2014). More recently, the other novel PEDV G2b variant with a 5-aa insertion (DTHPE) in the S gene was discovered in a PED-endemic farm in South Korea, thus suggesting that the virus undergoes continuous evolution in the field (Lee and Lee, 2017). The existence of these variants and their epidemics is unlikely at present but is still possible. Therefore, it is important to continue surveying hitherto-unidentified PEDV variants that may emerge locally or globally through genetic drift (e.g. point mutations) or genetic shift (e.g. recombination events) in order to prepare for the eventuality of future epizootics or panzootics. Control of the virus Biosecurity One of the most important measures for prevention and control of acute PED outbreaks is stringent biosecurity that fundamentally reduces the risk of the entrance of the virus into pig farms (especially fattening and farrowing units) by minimizing the introduction of any

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material or person that could be in contact with the virus. To accomplish this, all transport vehicles, materials associated with personnel including external visitors (hands, coveralls, and boots), and incoming fomites that could be contaminated with PEDV are thoroughly disinfected (Fig. 5.8). Although PEDV is inactivated by most virucidal disinfectants (Pospischil et al., 2002), PEDV RNA can still be detected by RT-PCR even after disinfection with a number of commercially available disinfectants (Bowman et al., 2015). Thus, it may be necessary to evaluate disinfectants in vivo or under various field conditions, particularly during the winter season, in order to select suitable disinfectant compositions and pertinent procedures. The following order of disinfection protocols is critical and recommended for pork producers attempting to sanitize transportation equipment or swine facilities that have moved or housed PEDV-positive animals: (i) proper cleaning using a high-pressure washer and warm water at temperatures over 70°C; (ii) disinfection with an appropriate disinfectant according to directions on the label; and (iii) overnight drying (Park and Lee, 2009; Park et al., 2011; Lee, 2015). Other biosecurity measures include restricting human traffic between fattening and farrowing units, and limiting contact between trailers or drivers and the farm interior during the loading process at the pig farm, or between drivers and the slaughter facilities during the unloading process at the collection point (Park and Lee, 2009; Park et al., 2011; Lowe et al., 2014). All newly arriving or replacement animals including gilts should be isolated for a certain period to monitor their health status (Park and Lee, 2009; Park et al., 2011). More importantly, when suspected clinical signs occur on pig farms, prompt notification followed by early diagnosis for PEDV by RT-PCR is also vital to minimize the spread of infection and to control PED. Clearly, personnel compliance is the key to successful application of such procedures. PEDV is a transboundary virus that seems to spread readily to neighbouring or distant countries, and even across continents. Because PED is not a World Organization for Animal Health reportable disease, countries may not properly implement quarantine inspection in potential sources or routes that mediate virus transmission between pork-producing nations. During large-scale, severe PED epidemics in adjacent or trading countries, quarantine (or international biosecurity) procedures should be adequately tightened with particular attention to any risk factors of global transmission in order to prevent the entrance of PEDV, as

Figure 5.8  Biosecurity and disinfection procedures for the prevention of PEDV spread. Biosecurity programmes must be implemented to anything or anyone sourced inside or outside to effectively minimize the risk of PEDV entry to herds, and to aid in reduction of viral spread between regions or nations.

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well as other emerging or re-emerging pathogens. Considering recent international spreads of global PEDV strains within Asian countries, between Asia and North America and to Europe, we may re-evaluate the importance of a quarantine policy against PEDV and other epizootic or zoonotic agents (Lee, 2015). Vaccines Passive immunization through sow vaccination is the most promising and effective strategy to control and eradicate PED. Because of an impermeable placenta caused by the absence of an immunoglobulin receptor, piglets are born agammaglobulinic and are highly susceptible to a variety of infectious agents (Langel et al., 2016). Therefore, newborn piglets rely exclusively on a transfer of maternal antibodies via colostrum and milk from immune dams for their protection. Despite the first epizootics in European countries, these countries have not faced financial threats to justify vaccine development. On the other hand, serious PED epizootics in Asian countries have been frequent and therefore served as an incentive to develop a plethora of PEDV vaccines. In China, CV777-attenuated or -inactivated vaccines have been routinely utilized against PED, but PED outbreaks in vaccinated herds challenged the effectiveness of the CV777-based vaccine (Li et al., 2012). The virulent Japanese PEDV strain 83P-5 was attenuated after 100 passages in Vero cells (Sato et al., 2011). Subsequently, the cell-adapted 83P-5 strain has been employed as an intramuscular (IM) live-attenuated vaccine (P-5V) in Japan, and it is also available in South Korea. The cell-culture adaptation method was also explored to attenuate two South Korean virulent PEDV strains, SM98-1 (93 passages) and DR-13 (100 passages) (Kweon et al., 1999; Song et al., 2007). The SM98-1 strain has been used as an IM live or killed vaccine, whereas DR-13 is available as an oral live vaccine. Although these attenuated or inactivated vaccines have been demonstrated to provide protection under experimental trials, their effectiveness in the field, as well as the pros and cons of their use, have been debated. The incomplete effectiveness of the aforementioned PEDV vaccines in China and South Korea might result from antigenic, genetic (variation between S proteins), and phylogenetic (G1 vs. G2) differences between vaccine and field epidemic strains (Lee et al., 2010, 2015; Lee and Lee, 2014; Oh et al., 2014; Kim et al., 2015). Therefore, G2b epidemic PEDV or related strains dominant in the field should be considered as seeds for developing the nextgeneration vaccines. The recombinant S1 protein derived from the field G2 PEDV isolate efficiently protected newborn piglets against PEDV, indicating its potential use as a subunit vaccine for PED prevention (Oh et al., 2014). The PED RNA vaccine was developed using the S gene from a G2b isolate in the RNA particle technology platform, based on a replication-deficient Venezuelan equine encephalitis virus packaging system in the USA (Mogler et al., 2014). Although the use of the PED RNA vaccine significantly increased the S-specific IgG antibodies in the colostrum, piglet mortality was barely reduced (Grenier et al., 2015). A breakthrough was achieved in the development of effective G2b-based vaccines with the isolation of PEDV strains phenotypically and genotypically identical to field strains responsible for global PED epidemics. A number of culturable PEDV strains associated with recent emergence were secured in the USA (Chen et al., 2014; Oka et al., 2014). On the basis of these G2b isolates, an inactivated PEDV vaccine has been manufactured and is currently available on the USA and South Korean markets. In South Korea, isolation and propagation of an epidemic virulent PEDV G2b strain in cell culture succeeded in our laboratory, and we are now investigating this isolate to spur the development of new effective

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and safe vaccines (Lee et al., 2015, 2017; Baek et al., 2016). Intriguingly, previous exposure of sows with ‘low-pathogenic’ G1b PEDV was shown to provide cross-protective lactogenic immunity in piglets challenged with ‘high-pathogenic’ G2b virus (Goede et al., 2015). However, sows receiving a parenteral vaccination developed a specific immune response that was not fully protective for their piglets, and this was reproducible in a young pig model (Crawford et al., 2016). These findings suggest that the route of vaccine administration may be one of the most momentous factors in stimulating the optimum mucosal immunity in sows, and subsequent transferring quality passive lactogenic immunity to suckling neonates for protection against PEDV. Although protection against the enteric disease is primarily dependent on the presence of secretory IgA antibodies in the intestinal mucosa, the vaccine efficacy might be associated with maintaining high levels of PEDV-specific neutralizing antibodies in the serum and colostrum of vaccinated sows (Park and Lee, 2009; Park et al., 2011; QIA, 2014). Song et al. (2016) suggest that colostrum and milk IgA and PEDV neutralizing antibody titres may be a correlate for protective immunity against PEDV. Other research shows that viral dose and the extent of viral replication in the gut of the gilt may contribute to production of sufficient levels of IgA and neutralizing antibodies in lactogenic secretions (Langel et al., 2016). Considering the issues mentioned above, there is a strong demand for the development of a new modified live virus (MLV) vaccine using the field-dominant strains to efficiently induce lactogenic immunity, and it is expected that advanced G2b-based MLV vaccines will soon be commercially ready for pig producers. However, there is still a gap in our understanding of the elements that influence induction of lactogenic immunity, which may include administration dose, vaccine strain, and the age or parity of the gilt/ sow, as well as other variables. Further research is needed to identify those factors, thereby ultimately improving vaccine regimens and overall herd immunity. Although it may be difficult to predict the efficacy of new vaccines in the field, it is important to bear in mind that vaccination will be the most valuable practical tool for prevention and/or control of PED, if its use is combined with stringent biosecurity/disinfection procedures and optimal farm and husbandry managements. Integrated and coordinated efforts among researchers, swine veterinarians, producers, swine industry specialists, producer associations, and authorities are required to achieve effective implementation of necessary PED prevention and control measures. Another crucial aspect in passive lactogenic immunity is to fruitfully supply adequate quantities of protective antibodies obtained from sow colostrum and milk to neonatal suckling piglets. Therefore, sanitation and health conditions of pregnant or lactating sows have to be monitored to eliminate potential factors that wane lactation performance, such as mastitis or agalactia, so that sows could constantly provide high-quality colostrum and milk to their litters (Lee, 2015). Suckling piglets are deprived of their source of maternal lactogenic immunity at weaning, and soon thereafter become vulnerable to PEDV. Following PEDV outbreaks, the virus may persist in susceptible animals or pigs that survived, leading to the circulation of the virus on the farm (endemic PED). Thus, active immunization of weaner to finisher pigs may be necessary for the control of endemic PEDV infections (Saif et al., 2012; Lee, 2015). Alternative immunoprophylactic and therapeutic strategies We could consider intentional exposure (feedback) of pregnant sows to the autogenous virus, during unexpected PED epidemics with high mortality rates, using watery faeces

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or minced intestines from infected neonatal piglets might be considered, which will artificially trigger rapid lactogenic immunity and, hopefully, shorten the outbreak period on the farm (Saif et al., 2012; Lee, 2015). However, there exist a number of drawbacks that should be contemplated before adopting this approach. Widespread dissemination of other viral pathogens, such as PRRSV or PCV2, that might be present in the intestinal or faecal contents, may emerge among sows or piglets, or within the farm ( Jung et al., 2006; Park et al., 2009). Since homogeneity and the amount of infectious PEDV are immeasurable in autogenous viral materials, sow immunity may not be sufficiently stimulated to a quality satisfactory for offspring protection. Furthermore, infectious viruses derived from artificial exposure of sows will be shed in faeces or oral fluids, which, in turn, could be a potential source for PEDV transmission within the contaminated establishment and between facilities and farms (Lee, 2015). Artificial passive immunization by oral administration of specific antibodies represents an attractive approach against enteric pathogens such as PEDV. The immunoprophylactic effects of cow colostrum and chicken egg yolk immunoglobulin against PEDV or the S1 gene have been shown to protect neonatal piglets following exposure to a challenge (Kweon et al., 2000; Shibata et al., 2001; Lee et al., 2015). Pharmacological, biological, or natural agents that decelerate epithelial cell renewal by stimulating proliferation or reorganization of crypt stem cells could be potential therapeutic targets to reduce PEDV-associated mortality from dehydration resulting from severe villous atrophy ( Jung et al., 2015). The epidermal growth factor was found to stimulate proliferation of intestinal crypt epithelial cells and relieve PEDV-induced atrophic enteritis, indicating its potential as a therapeutic option ( Jung et al., 2008). Broad-spectrum antiviral drugs or molecules, such as ribavirin, which suppress PEDV infection in vitro, are of interest for their practical uses to treat PED (Kim and Lee, 2013). Chemical inhibitors as well as compounds from medicinal plants or natural sources, which act on extracellular targets such as cholesterol-dependent virus entry ( Jeon and Lee, 2017) or intracellular factors including apoptosis or MAPK signalling pathways (Kim and Lee, 2014, 2015; Lee et al., 2016), could be novel therapeutic candidates to diminish PEDV-associated symptoms and mortality. Additionally, nutritional supplements that reduce stress and enhance resistance to the disease may be useful for PED control in neonates. Acknowledgements I thank Sunhee Lee, a graduate student at Kyungpook National University, for her assistance in the preparation of references and figures. This was supported by Bio-industry Technology Development Program through the Korea Institute of Planning and Evaluation for Technology in Food, Agriculture, Forestry and Fisheries (iPET) funded by the Ministry of Agriculture, Food and Rural Affairs (315021-04). References

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Porcine Parvovirus André Felipe Streck1* and Uwe Truyen2

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1

Veterinary Diagnostic Laboratory, Faculty of Veterinary Medicine, University of Caxias do Sul, Caxias do Sul, Brazil. 2 Institute for Animal Hygiene and Veterinary Public Health, Faculty of Veterinary Medicine, University of Leipzig, Leipzig, Germany. *Correspondence: [email protected] https://doi.org/10.21775/9781910190913.06

Abstract Porcine parvovirus (PPV) is considered the main cause of reproductive disorders in pigs, which are summarized under the acronym SMEDI (stillbirth, mummification, embryonic death, and infertility). In this chapter the biology of the virus and its structure, pathogenic potential and strain variation, as well as the disease induced by the virus, are described. Known aspects of pathogenesis, diagnosis and prevention, particularly by vaccination, are summarized. Furthermore, in recent years ‘new’ parvoviruses (PPV2 to 7) have been described in pigs. They have been detected in pigs from various parts of the world and their association with clinical signs or disease will be discussed. Introduction Porcine parvovirus (PPV) is a small non-enveloped virus considered to be one of the major causes of reproductive failure in swine worldwide. Historically, the reproductive losses in commercial swine herds were high in the 1960s, and, at that time, they were associated with environmental, nutritional, genetic and toxicological problems (Lawson, 1961; Rasbech, 1969). The first evidence of porcine parvovirus was obtained in primary cell cultures from porcine kidney and testicle used to cultivate hog cholera virus, where persistent contaminant small particles (22–23 nm size) were found (Mayr and Mahnel, 1964). These particles were similar to the Kilham rat virus (a parvovirus) (Mahnel, 1965). Due to the replication ability of the virus in cell lines from swine, it was possible to isolate and classified as a porcine parvovirus (Siegl 1976). The occurrence of PPV in pigs was first described by Cartwright and Huck (1967) and was associated with abortions. In the following years, PPV was identified as the main agent of recurring oestrus, abortion and the delivery of mummified or stillborn fetuses, commonly described by the acronym SMEDI (stillbirth, mummification, embryonic death and infertility). The virus is considered to be endemic in most areas of the world and can be found in all pig herd categories. Reproductive losses are typically low in vaccinated herds, but PPV

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can cause devastating abortion storms in unvaccinated herds, in situations in which the vaccine was administered incorrectly or in the emergence of new antigenic types (Truyen and Streck, 2012). Taxonomically, according to the last release of the International Committee on Taxonomy of Viruses (ICTV), the virus is a member of the family Parvoviridae, Parvovirinae subfamily and Ungulate protoparvovirus 1 species. Molecular biology of the virus The genome all of parvoviruses represents a single stranded (ss) DNA molecule of about 5 kb. In both terminal sequences of the virus, a complex palindromic hairpin structure of about 120–200 bases is located as requirement for DNA replication. The genome of PPV encodes four proteins and uses alternative splicing to extend the coding capacity. Two nonstructural proteins, NS1 and NS2, operate in the replication of the virus, particularly for DNA replication. Two structural proteins (VP1 and VP2) are transcribed and translated from the parvovirus genome. The smaller protein (VP2) is produced by splicing from the same RNA template as the larger protein (VP1), therefore VP1 has 729 amino acid residues, of which 120 form an amino-terminal unique portion (absent in VP2). The third structural protein, VP3, is a post-translational modification product of VP2 (Simpson et al., 2002; Cotmore and Tattersall, 2006). Additionally, a late non-structural protein (SAT) expressed from the same mRNA as VP2 is found seven nucleotides downstream of the VP2 start codon (Zádori et al., 2005). As shown in Fig. 6.1, PPV capsid is a spherical shell (with about 28 nm in diameter consisting) of 60 identical copies of these viral proteins arranged in an icosahedral symmetry (Chapman and Rosmann, 1993). The identical copies (called subunits) are built from about 90% of VP2 and 10% of VP1 molecules. (Simpson et al., 2002). The subunit consists of eight antiparallel β-strands, a common structure for viral capsids, together with one α-helix and four loops (Chapman and Rossmann, 1993). On the surface, a projection at the 3-fold axis, a depression or canyon around the 5-fold axis and a dimple on the 2-fold axis of symmetry

Figure 6.1  The capsid structure of the PPV. Left figure: Surface representations of the capsid calculated from X-ray coordinates with a low pass filter at 17 Å in a temperature factor of 500 Å. Right figure: 3D model of the PPV VP2 proteins using the cartoon technique, with a rocket (α-helix) and arrows (β-strands) representing the secondary structure. The image was generated with the software Cn3D version 4.1. The coordinates of both figures were retrieved from the NCBI Structure Database (http://www.ncbi.nlm.nih.gov/Structure/index.shtml). Accession number: 1K3V (Simpson et al., 2002). The neighbouring 5- (in blue), 3- (in red) and 2-fold (in green) axes of the capsid subunit are shown.

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can be observed. The 2- and 3-fold axes consist mainly of amino acids located in the subunit loops (Simpson et al., 2002). Cellular biology of the virus (including cell tropism, virus–cell interactions, cellular death, etc.) The primary replication of the PPV occur in lymphoid tissues. After that, the virus is distributed systemically via viraemia (Paul et al., 1980). It is still not understood how the PPV crosses this placental barrier and reaches the fetus, since six tissue layers completely separate the sow and the fetal blood circulation and these cells are closely connected, not allowing the passage of even small molecules as antibodies, (Mengeling, 2000). Probably, as the virus is able to remain infectious after phagocytosis by macrophages, it can cross the porcine epitheliochorial placenta using macrophages to infect the fetus (Paul et al., 1979). After fetal infection, the PPV has an environment particularly susceptible to infection and to replication due to the high mitotic activities present in the fetuses’ tissues. PPV entrance in a cell is still unclear, but include clathrin-mediated endocytosis, or micropinocytosis transportation mediated by the endosomal pathway (Boisvert et al., 2010). Endosomal trafficking and acidification are essential for PPV to enter in the nucleus (Boisvert et al., 2010), due to reversible modifications of the capsid allowing the virus to escape from the endosome (Vihinen-Ranta et al., 2002; Farr et al., 2005). After the virus arrives in the nucleus, PPV replicates using the cell’s own replication mechanism. The virus replicates in cells in replication phase (S) using the cellular DNA polymerase. This explains the requirement for cells with a higher replication index (Rhode, 1973). PPV replication decreased mitochondrial membrane potential and the subsequent oxidative damage also leads to the release of cellular toxic proteins such as cytochrome c from the mitochondria to the cytosol, triggering apoptosis and causing cell death and tissue damage in viral diseases (Zhao et al., 2016). The virulence properties of the PPV appear to be related to the viral protein gene. In vitro studies using recombinant viruses derived from pathogenic (Kresse) and non-pathogenic (NADL-2) PPVs showed that single amino acids in the capsid protein affected an isolate’s capacity to replicate in certain cell lines. Furthermore, a comparison between Kresse and NADL-2 genomes showed that the non-coding regions were nearly identical. For the nonstructural gene region (NS1/NS2), all differences found are silent (synonymous), while in the structural genes (VP1/VP2) six of eight differences led amino acid substitutions (nonsynonymous). Among the VP2 amino acids, five changes were consistent with comparisons in field isolates (I-215-T, D-378-G, H-383-Q, S-436-P, and R-565-K) and three of these (D-378-G, H-383-Q, and S-436-P) were considered responsible for differences in tissue tropism (Bergeron et al. 1996). The amino acid position 436 is located right in the 3-fold spike centre of the capsid subunit and the amino acid position 215 at its base. This location in the 3-fold spike has been considered to be an important antigenic surface region in various parvoviruses (Chapman and Rossmann 1993). In the recent viral sequences obtained, it was found that substitutions were mainly located in the capsid surface, therefore influencing the receptor binding and/or antigenicity (Streck et al., 2015a). This could be evident observing the neutralization activities of sera raised against two recent German field isolates by the experimental infection of pregnant sows at day 40 of gestation. The post-infection sera of these sows were tested for their homologous and heterologous neutralization activities. All the antisera demonstrated a

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high neutralization activity against the homologous viruses, but lower activity against the prevalent field strain in Germany (27a) (Zeeuw et al., 2007). Thereafter, studies observed strains closely related to the 27a strain in several countries and it was estimated that the main divergences between the new isolates were probably introduced in the last 10–30 years (Cadar et al., 2011; Streck et al., 2011). Those findings lead to the hypothesis that the emergence and predominance of ‘27a-like’ strains could be a viral adaptation to the largely used vaccines resulting in ‘escape mutants’ (Fig. 6.2). Clinical and pathological observations The major clinical sign of PPV infections is maternal reproductive failure reproductive failures. Diarrhoea and skin lesions were also linked to PPV; however, the aetiological role of the virus remains to be fully established (Brown et al., 1980; Dea et al., 1985; Duhamel et al., 1991). Subclinically, a moderate and transient lymphopenia, independent of sex and age, can be observed between 5 and 10 days after initial infection ( Joo et al., 1976; Mengeling and Cutlip, 1976; Zeeuw et al., 2007). Pathological sequela caused by PPV is related mainly to the gestational period in which infection occurs. At the gestation beginning, the conceptus is protected by the zona pellucida and is not susceptible to infection. Thereafter in the stage of embryo (until approximately day

Figure 6.2  PPV sequences phylogenetic tree inferred using the Maximum Likelihood method based on the Hasegawa-Kishino-Yano model. A discrete Gamma distribution was used to model evolutionary rate differences among sites (5 categories). The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Evolutionary analyses were conducted in MEGA7 (Kumer et al., 2016). The reference strains NADL-2, Kresse and 27a are indicated by black dots.

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35 of gestation) an infection with PPV results in embryonic death and maternal resorption of fetal tissues. From the 35th day of gestation on, fetal organogenesis is mainly complete and ossification of the fetal skeleton begins. PPV infection after this time typically results in fetal death followed by mummification. Finally, after the 70th gestation day, the fetus is able to build an effective immune response and eliminate the virus and fetal infection becomes mainly subclinical. In this last case, the piglet is born with anti-PPV antibodies (Bachmann et al., 1975; Joo et al., 1977; Lenghaus et al., 1978; Mengeling et al., 2000). Lesions are only evident in embryos or fetuses. Even experimental PPV inoculation in boars, gilts, and sows does not produce macroscopic lesions (Bachmann et al., 1975; Mengeling and Cutlip, 1976; Lenghaus et al., 1978; Mengeling, 1978; Thacker et al., 1987). Macroscopically, the embryonic death followed by resorption of fluids and soft tissues is the most common sequel to PPV infection. The lesions include a variable degree of congestion, oedema and haemorrhage with accumulation of serosanguinous fluids in body cavities. After the death of the fetus, a discoloration of the skin resulting from the bleeding occurs, giving the fetus a dark tonality. Finally, progressive dehydration of tissues leads to the mummification (Fig. 6.3). The placenta can be dehydrated and brown to grey in colour and the extrafetal fluid volume reduced ( Joo et al., 1977; Lenghaus et al., 1978). Microscopic lesions in females include focal accumulation of mononuclear cells adjacent to the endometrium and in deeper layers of the lamina propria and a marked perivascular cuffing of plasma cells and lymphocytes in the brains, spinal cord, and choroid of the eye (Hogg et al., 1977). Sows also had uterine lesions that included extensive cuffing of mononuclear cells around myometrial and endometrial vessels (Lenghaus et al., 1978). In the fetus, histopathological changes tend to be widespread in the different tissues and the major lesions represent necrosis of cells in developing organ systems ( Joo et al., 1977; Lenghaus et al., 1978). Haemorrhages are present in subcutaneous tissues and muscle masses. Necrosis and mineralization are common in the lungs, kidneys, and skeletal muscle,

A

B

Figure 6.3  Litters of inoculated pregnant sows [with 27a (A) and NADL-2 strains (B)] at the 90th gestational day displaying distinct levels of lesions. The fetuses’ position corresponds to their position in the uterus (the most cervical-positioned fetuses at the top) (Zeeuw et al., 2007).

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particularly in the liver and heart (Lenghaus et al., 1978). After the fetuses become immunocompetent, microscopic lesions are primarily endometrial hypertrophy and mononuclear cell infiltration (Hogg et al., 1977; Joo et al., 1977). Meningoencephalitis characterized by perivascular cuffing with proliferating adventitial cells, histiocytes, and a few plasma cells in the grey and white matter of the cerebrum and leptomeninges were also seen in PPVinfected live fetuses delivered late in gestation or in stillborn piglets (Narita et al., 1975; Hogg et al., 1977; Joo et al., 1977). Diagnostic procedures Porcine parvovirus infection may not cause abortions and does not cause clinical signs in adults (Mengeling and Cutlip, 1975). PPV can be strongly considered when females return to oestrus with no apparent reasons or delays in parturition with increased numbers of mummified fetuses and smaller litters, especially in first or second parity females. For these clinical signs, the differential diagnosis should also include Aujeszky’s disease, brucellosis, leptospirosis, porcine reproductive and respiratory syndrome (PRRS), toxoplasmosis, nonspecific bacterial uterine infection, and other metabolic and genetic problems. The material to be submitted to the laboratory to perform the diagnostic should include mummified fetuses and fetal remains. Detection of viral antigen in fetal tissues by immunofluorescence (IF; Fig. 6.4) was a reliable procedure for the diagnosis of PPV (Mengeling and Cutlip, 1975). In the past, the virus detection and titration can be performed by the hemagglutination technique (HA), based on the hemagglutinating activity of PPV against erythrocytes of certain species, e.g. chickens, humans, guinea pigs ( Joo et al., 1976). Today, since the virus has a high replication efficiency in renal or testicular swine cells, virus propagation in cell-lines such as ESK (embryonic swine kidney), PK-15 (pig kidney), SK6 (swine kidney), STE (swine testicular epithelioid) and SPEV (swine embryo kidney) is used. Once in these cells, the PPV replication usually causes cytopathic effects, including granulations, irregular shape, slow replication, intra-nuclear inclusions, pyknotic nucleus and consequently cell death (Cartwright et al., 1969; Mengeling, 1972). Due to possible similar cytopathic effects of other viruses or enzymatic effects, virus isolation and titration is often associated with immunofluorescence microscopy (Cartwright et al., 1971; Johnson, 1973).

Figure 6.4  Indirect immunofluorescence of PK-15 cells infected with PPV. Positive nuclear fluorescence (green) is evidenced five days post infection. Magnification of 400x.

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More recently, nucleic acid-based techniques can be used for viral detection in clinical samples with a better sensitivity that the other direct techniques. The polymerase chain reaction (PCR) is the most useful technique for the detection of PPV in fetal tissues, semen, and other samples. Numerous PCR protocols (and quantitative PCR methods) have been described (Molitor et al., 1991; Soares et al., 1999; Wilhelm et al., 2006; Chen et al., 2009; Streck et al., 2015b; Yang et al., 2016). These methods are considered to possess higher diagnostic sensitivity and specificity than hemagglutination or virus isolation and are better suited for the detection of PPV in autolysed tissues. However, the likelihood of successful recovery of virus or nucleic acid will depend on the condition of fetal tissues at the time of collection. Alternatively, serology may be useful for the diagnosis of PPV when fetal tissues are not available or sample is autolysed; however, the high prevalence of PPV in populations often present challenges to the interpretation of results. For these reasons, paired serum samples should be evaluated. As standard method, the haemagglutination inhibition (HI) assay is frequently used for the quantification of PPV specific antibodies. Usually, the serum to be assayed in the HI test is usually pretreated by heat inactivation (56°C, for 30 minutes) followed by adsorption with erythrocytes and kaolin (to remove or reduce non-specific inhibitors of haemagglutination) (Mengeling, 1972; Morimoto et al., 1972). Importantly, HI results may be affected by incubation temperature and the source of erythrocytes. Another technique, the enzyme-linked immunosorbent assay (ELISA), can be standardized more easily and is suited to automatization. Moreover, serum does not have to be pre-treated before testing in the ELISA (Hohdatsu et al., 1988; Westenbrink et al., 1989). The ELISA can potentially differentiate vaccinated animals from animals having been infected with PPV. As the currently used inactivated vaccines induce antibodies only against VP proteins and not against the NS proteins, ELISAs that differentiate these two proteins could identify antibodies raised from a natural infection (Madsen et al., 1997; Qing et al., 2006). Epidemiology PPV is considered to be endemic in most of the world. The virus can be found in all pig herd categories, including in boars and fattening pigs. The epidemiology of porcine parvovirus is primarily marked by the high stability of the virus in the environment. That is, PPV can remain infectious for months, and contaminated instruments or stables may therefore be a constant source of infection. The virus can be transported between herds via fomites, for example the clothes, boots, equipment and clothing of farmers from one herd to another. It is also speculated that infected boars can introduce the virus into new herds. There are numerous reports on PPV in the semen of naturally infected boars (Cartwright and Huck, 1967; Ruckerbauer et al., 1978); however, whether PPV is shed in the semen of infected boars or whether PPV in semen represents only a contamination is still unresolved. Importantly, introduction of PPV into a herd does not cause immediate problems if sows are immune through vaccination or natural exposure. Disease occur when there is circulation in a population with new mutants that cannot be completely prevented by vaccination or a lack of vaccination. Stability of PPV can be evidenced by its resistance to inactivation by ethanol (70%), quaternary ammonium (0.05%) and low concentrations of sodium hypochlorite (2500 ppm) and peracetic acid (0.2%). The virus is also heat stable and may resist dry (but not moist)

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heat at 90°C. The inactivation of PPV should be performed by aldehyde-based disinfectants, higher concentrations of sodium hypochlorite (25,000 ppm) and hydrogen peroxide (7.5%) (Eterpi et al., 2009). Control of the virus (including vaccine and antiviral research) There is no specific treatment for parvovirosis. General management measures aimed at promoting good health status of the herd should be adopted (Mengeling, 1999). A more practical goal in commercial herds is to maintain herd immunity against PPV. Before the advent of vaccination, intentional infection of gilts to PPV by exposure to virus-contaminated tissues from affected litters before the first breeding was used to control parvovirus. Approaches like this type are unreliable and dangerous because they can result in the dissemination of other pathogens in the population, for example, classical swine fever virus. The first vaccines developed during the 1970s were made with inactivated virus (Suzuki and Fujisaki, 1976, Joo and Johnson, 1977, Mengeling et al., 1979). A few years later, regular vaccination of breeding sows with these vaccines became a worldwide practice. Currently, PPV vaccines represent cell culture derived virus (usually the non-pathogenic NADL-2 strain) which is chemically inactivated (by formalin, beta-propiolactone or binary ethyleneimine), mixed with oil or aluminium hydroxide as adjuvants and administered parenterally. The use of these vaccines induces antibody titres that can reduce clinical manifestations, but cannot prevent infection ( Józwik et al., 2009, Foerster et al., 2016). The vaccination schedules can be adjusted to the animal category. Gilts usually receive the first dose at the age of 170–180 days, or 30 days before insemination. The second dose is usually administered 15 days later. Sows are boosted usually 10–15 days after each farrowing. Boars can be vaccinated as well and boosted yearly. Modified-live virus vaccines (MLV) could be an alternative for PPV. The use of parvovirus MLVs in carnivores induce a long-lasting immune response that provides protection for several years. For PPV, the few reports on live-vaccines observed transplacental transmission was prevented, but viraemia and shedding of the vaccine strain after vaccination was common (Paul and Mengeling, 1980, 1984). However, the attempts were mainly based on NADL-2 virus as the vaccine virus. To our knowledge, there are no reports of vaccines based on a modified strain. Furthermore, the occurrence and distribution of new antigenic types have to be watched very closely. It was observed that new mutations in the capsid protein can modify antigenic properties, and may reduce the cross neutralization ability of sera raised against strains commonly used in commercial vaccines (Zeeuw et al., 2007; Streck et al., 2013, 2015a). New vaccines that induce a long-lasting immunity and cover all the dominant virus strains circulating in pig populations are needed to overcome currently observed putative vaccination failures. The emergence of novel parvoviruses In other animal species, such as humans and canines, parvoviruses are represented by viruses classified in two or more genera. In pigs, only in the genus protoparvovirus a parvovirus, commonly known as porcine parvovirus (PPV) was established, however, in the last decade, with the development of novel molecular techniques several new (proto)parvoviruses have

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been identified. The first newly identified parvovirus was identified in sera during a hepatitis E virus investigation in Myanmar (Hijisaka et al., 2001), originally termed as porcine parvovirus 2 (PPV2). More recently, a high prevalence of this virus could be obtained in lung samples. Viral positivity could also be detected in faecal samples obtained from pigs of different age groups, and in sera or thoracic fluids obtained from neonatal pigs (Xiao et al., 2013). In 2008, a virus closely related to the human parvovirus 4 was found in slaughtered pigs in Hong Kong (Lau et al., 2008). Currently, several sequences with high homology (> 98% DNA similarity) exist in the DNA databanks under different names, including porcine hokovirus, PARV4-like and porcine parvovirus 4. The virus has already been detected worldwide (Adlhoch et al., 2010; Cadar et al., 2011; Pan et al., 2012; Xiao et al., 2012) and its usually nominated porcine parvovirus 3 (PPV3). In 2010, another parvovirus was found in the USA in porcine circovirus-associated disease-affected pigs and designated as porcine parvovirus 4 (PPV4) (Cheung et al., 2010). After that, PPV4 was already detected in several continents (Huang et al., 2011, Zhang et al., 2011; Cságola et al., 2012; Cadar et al., 2013; Ndze et al., 2013). Thereafter, during an initial investigation of the PPV4 prevalence in U.S. pigs, a novel PPV, named PPV5, was identified and showed the closest relationship to PPV4 (Fig. 6.5) (Xiau et al., 2013). Another closest related parvovirus to PP4 and PPV5 was initially identified from aborted pig fetuses in China and termed PP6 (Ni et al., 2014). Finally, in pigs with postweaning multisystemic wasting syndrome (PMWS), a porcine bocavirus (PBoV, classified in the genus Bocaparvovirus) was identified in lymph nodes (Blomström et al., 2009) and subsequently other novel bocaviruses named PBoV2, PBoV3,

Figure 6.5  Phylogenetic tree of the porcine parvoviruses sequences inferred using the using the Maximum Likelihood method. The confidence was measured using the bootstrap method inferred from 1000 replicates and it is displayed with colour gradient (1 = red; 0 = blue). A discrete Gamma distribution was used to model evolutionary rate differences among sites (5 categories). The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. Evolutionary analyses were conducted in MEGA7 (Kumer et al., 2016).

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Table 6.1 The formerly used nomenclature for the species and the current classification based upon the International Committee on Taxonomy of Viruses (ICTV) Virus

ICTV current classification

Porcine parvovirus 1

Ungulate protoparvovirus 1

Porcine parvovirus 2 Porcine parvovirus 3, porcine hokovirus, PARV4-like Porcine Cnvirus

Not yet classified Ungulate tetraparvovirus 2 Ungulate tetraparvovirus 3

Porcine parvovirus 4

Ungulate copiparvovirus 2

Porcine parvovirus 5

Not yet classified

Porcine parvovirus 6

Not yet classified

Porcine bocaviruses

Porcine bocaparvovirus 2, 3, 4 and 5

Table adapted from Streck et al. (2015a).

PBoV4, 6V and 7V followed (Cheng et al., 2010; Zhai et al., 2010; Zeng et al., 2011; Yang et al., 2012). Despite the epidemiological studies, any possible association with clinical signs of these newly identified PPV or PBoV remains unclear. Although, they may possess a potential public health interest due to their genetic similarity with human parvoviruses. The current classification based upon the International Committee on Taxonomy of Viruses for these novel viruses is available in Table 6.1. References

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Porcine Reproductive and Respiratory Syndrome Virus Alexander N. Zakhartchouk1*, Sujit K. Pujhari2 and John C.S. Harding3

7

1Vaccine and Infectious Disease Organization – International Vaccine Center (VIDO-InterVac),

University of Saskatchewan, Saskatoon, SK, Canada.

2Department of Entomology, Center for Infectious Disease Dynamics, and Huck Institutes of the

Life Sciences Millennium Science Complex, Pennsylvania State University, PA, USA.

3Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK, Canada.

*Correspondence: [email protected] This paper was published with the permission of the Director of VIDO-InterVac, journal series no. 812 https://doi.org/10.21775/9781910190913.07

Abstract Porcine reproductive and respiratory syndrome virus (PRRSV) is a swine arterivirus responsible for reproductive failure and respiratory problems in sows and piglets, respectively. It is one of the most economically devastating diseases of pigs and continues to result in major health challenges in the swine industry worldwide. The virus is continuously evolving and emerges episodically in different regions of the world with increased virulence. In this chapter, we briefly summarize the current understanding of PRRSV from the perspectives of the virus molecular biology, virus–host cell interactions, pathogenesis, diagnostic procedures and epidemiology. We also provide an overview of currently available vaccines and a novel vaccine development. History PRRSV is a relatively new virus, clinically reported first among swine populations in the late 1980s in the USA. Almost a decade later PRRSV re-emerged in Europe with symptoms previously unreported. For many years, until the etiological agent was identified, the disease resulting from PRRSV infection was colloquially referred to as ‘mystery swine disease’ (Hill, 1990; Reotutar, 1989). The virus was first isolated at the Central Veterinary Institute in the Netherlands in 1991, and shortly thereafter in 1992 in the USA and was designated Lelystad virus and swine infertility and respiratory syndrome (SIRS) virus (BIAH-001), respectively (Wensvoort et al., 1991; Collins et al., 1992). The virus is now commonly referred as porcine reproductive and respiratory syndrome (PRRS) virus. Molecular and serological data shows

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evidence for the presence of this virus in Europe and Canada as early as 1979 (Carman et al., 1995). It has since become a major problem to the swine industry worldwide, with estimated annual costs of 664 million U.S. dollars in the United States alone (Holtkamp et al., 2013). Classification PRRSV is categorized as an arterivirus of the family Arteriviridae along with simian haemorrhagic fever virus (SHFV) of monkeys, lactate-dehydrogenase elevating virus (LDV) of mice, equine arteritis virus (EAV) of horses, and newly recognized wobbly possum disease virus (WPDV) of free-ranging Australian brushtail possums (Baker, 2012; Plagemann and Moennig, 1992). Members of the family Arteriviridae are related to Coronaviridae, Roniviridae and Mesoniviradae. The former two are pathogens for vertebrate hosts, and latter two are of fish and insect species, respectively. Nested subgenomic (sg) messenger RNA is one of the prominent features of the genome transcription strategy by this group of viruses. This unique feature elevated them to a new order Nidovirales (nidus is the Latin for nest). Further, based upon their genome size, the order Nidovirales can be divided into three clades: the large nidoviruses (26.3–31.7 kb), which includes the Coronaviridae and Roniviridae; the medium nidoviruses (20 kb), which includes Mesoniviradae; and the small nidoviruses (12.7–15.7 kb), which includes Arteriviridae. Both European and North American PRRSV isolates cause similar clinical symptoms; however, they now represent two distinct viral species: PRRSV-1 (formerly European genotype 1) and PRRSV-2 (formerly North American genotype 2) whose genomes diverge by approximately 40% (Nelson et al., 1993; Nelsen et al., 1999). The European and North American PRRSV prototypes virus isolates are Lelystad virus and VR-2332, respectively. Each species contains subtypes and strains, which are genetically diverse and vary in virulence and pathogenicity. EAV is the best-characterized arterivirus, although recent studies have increasingly been focused on PRRSV due to its economic importance. Molecular biology of the virus Morphology of the capsid PRRSV is roughly spherical or oval in shape as seen under the electron microscope. The virus particles measure between 50 to 74 nm in diameter, with a median value of 54 nm. It has a mostly smooth, featureless surface with a few protrusions of membrane glycoproteins. Among the positive sense RNA viruses, members of the Arterivirus genus have the greatest number of structural proteins (nine for PRRSV) composing their envelope. Viral RNA and nucleocapsid protein forms a pleomorphic core that measures approximately 39 nm in diameter. It is separated from the outer envelope by 2–3 nm gap (Spilman et al., 2009). The details of structural arrangement of virus nucleocapsid are discussed below. Physiochemical property The buoyant density of PRRSV is 1.18 to 1.22 g/cm3 in sucrose, and the sedimentation coefficient of 214s–230s (Yuan et al., 2004). PRRSV is stable and can be stored at –70°C for long periods of time but loses it stability and infectivity at higher temperatures (Zimmerman et

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al., 2010). It is also highly unstable in solutions containing a low concentration of non-ionic detergents. The virus is fairly stable between pH 6.0 and 7.5, but is rapidly inactivated by high or low pH. Viral genome structure The virus has a single-stranded, non-segmented, positive sense RNA genome that is polyadenylated at the 3′-end and protected by 7-methylguanine cap at 5′-end. The viral RNA is polycistronic and infectious in nature. The genome of PRRSV-2 (15.5 kb) is a few kb longer than PRRSV-1 (15.1 kb). PRRSV shows close similarity in genomic organization with EAV and LDV. EAV is the prototype of Arterivirus genus, with a genome length of about 12 kb. LDV, the closest relative of PRRSV, has a genome length of ~14 kb. The PRRSV genome encodes for at least ten open reading frames (ORFs) flanked by 5′ and 3′ untranslated regions (UTRs) (Fig. 7.1). The 5′ UTRs are 217–222 nucleotides (nt) in length for PRRSV-1 and 188–191 nt for PRRSV-2; the 3′ UTRs are approximately 150 nt (114 nt in PRRSV-1 and 148 nt in PRRSV-2) excluding the polyadenylation site (Verheije et al., 2002; Beerens and Snijder, 2007). The non-structural replicase genes encode two large ORFs, 1a and 1b, which occupy nearly three fourths of the 5′-end of the genome. Using a ribosomal frameshift event, ORF1a and ORF1b produce two large precursor polyproteins, pp1a and pp1ab, and two small

Figure 7.1  PRRSV structure and genome organization. (A) Schematic presentation of the PRRS virus particle. The roughly spherical virus particle has an outer lipid bilayer envelope derived from the host cell. Viral structural proteins (except N protein) remain embedded in the bilayer membrane with trifling projections, which makes the virus surface almost smooth. GP4 and GP2 mediate the interactions with the other minor viral membrane proteins and interact with the CD163 receptor on the host cell mediating virus entry. GP5 and M, also found on the virus envelope, form a heterodimeric complex and facilitates the virus assembly process. The envelope encloses the inner hollow nucleocapsid. Viral RNA genome and N protein form the nucleocapsid. (B) PRRSV has ~15 kb positive sense RNA genome. Almost three quarters of the genome at 5′-end encodes for two polyproteins (pp1a, and pp1ab) that are expressed using programmed ribosomal frameshifting (PRF) and subsequently cleaved into 14 non-structural proteins. In addition, two more proteins (nsp2TF and nsp2N) are produced using PRF. The rest of the genome encodes for eight structural proteins that are translated from a 3′-co-terminal nested set of sgRNA (sg mRNA2–7).

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proteins, nsp2N and nsp2 transframe fusion (TF) (Fang et al., 2012; Snijder et al., 2013). The polyproteins are subsequently processed into 16 non-structural proteins that include proteases, helicase, and the RNA-dependent RNA polymerase (Table 7.1). The expression of ORF1b is mediated by a –1 programmed ribosomal frameshift (PRF) just before the stop codon for ORF1 which allows production of the pp1ab protein. Another –1 PRF occurs somewhere in the middle of ORF1a and produces a truncated version of pp1a known as nsp2N. In addition, a –2 PRF has been recently demonstrated, which produces a short TF protein similar to nsp2N but with an added transmembrane domain. The ribosomal slippage sequences mapping to the genomic positions 3889 nt and 7695 nt of VR-2332 are responsible for these PRFs. The remainder of the genome encodes for eight relatively small genes. All structural genes are translated from subgenomic (sg) mRNAs, a set of transcripts that have same 5′-leader sequence derived from the original genome (a feature of the order Nidovirales) (van Marle et al., 1999). ORF3 to ORF5 encode for glycoprotein (GP) 3 to 5, respectively. ORF2 produces GP2 protein. ORF6 encodes for membrane (M) protein, and ORF7 encodes for nucleocapsid (N) protein. ORF2 and ORF5 are functionally bicistronic, and they produce two additional proteins 2b (E) and 5a (Wu et al., 2001; Johnson et al., 2011). Except for the N protein, the structural proteins are part of the virus envelope. Together GP2, GP3, and GP4 form a trimeric complex resulting in the minor glycoprotein complex which functions in the viral entry process (Wissink et al., 2005). GP5 along with M form a disulfide-linked heterodimer and together constitute the major glycoprotein complex on the virion (Van Breedam et al., 2010). Genome transcription and replication The order Nidovirales, to which PRRSV belongs, has the most complicated genome transcription and replication process among the known positive sense RNA viruses. A large polycistronic RNA genome, use –1 and –2 PRF to accommodate additional functional protein sequences, and transcription of a nested set of 3′-end co-terminal sg mRNAs through a discontinuous transcription strategy augments several layers of complexity (Gorbalenya et al., 2006; van Hemert et al., 2008). Like other positive sense RNA viruses, PRRSV replicates in the cytoplasm. Of note, the two viral proteins nsp1 and N shuttle between the nucleus and cytoplasm (Rowland et al., 2003). The functional significance of this activity is yet to be understood. Upon PRRSV entry into a susceptible cell, the viral genome is released into the cytoplasm and acts as the mRNA (Delrue et al., 2010). It produces the two large nonstructural polyproteins pp1a and pp1ab, which encode for proteins that are necessary for the functional viral replication complex. Further, the same genomic strand is used to generate an anti-genome (negative strand RNA), which is the template for genome replication (Fig. 7.2) (Kappes and Faaberg, 2015). The early viral non-structural proteins assemble into an enzyme complex called replication and transcription complex (RTC), which associates with virus-induced double membrane vesicles (DMV) localized to the perinuclear space of the infected cell, where viral RNA synthesis occurs. The DMVs are derived from endoplasmic reticulum with the help of putative viral membrane spanning proteins nsp2, nsp3 and nsp5 (Pedersen et al., 1999). Besides viral proteins, cellular proteins may also play an important role in this process of membrane rearrangement, which is still a matter of investigation. The conserved ORF1b encodes the core replicative machinery: the RNA-dependent RNA polymerase

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Table 7.1 PRRSV proteins and their functions Number of amino acids Proteins

ORFs

Type I

Type 2

Function

180

180

Protease (PLP1α); zinc-finger protein; regulator of sg mRNA synthesis, IFN inhibition; distributes in the both nucleus and cytoplasm.

nsp1β

205

203

Protease (PLP1β); regulates the generation of nsp2TF; IFN inhibition; suppresses host mRNA export from the nucleus to the cytoplasm.

nsp2

1060

1196

Protease (PLP2) with deubiquitinating activity; DMV formation; IFN inhibition; incorporated into virion.

Non-structural proteins nsp 1α

ORF1a

nsp3

230

230

Integral membrane protein; DMV formation.

nsp4

203

204

Main protease (3C-like serine protease); IFN inhibition; apoptosis inducer.

nsp5

170

170

Integral membrane protein; DMV formation.

nsp6

16

16

?

nsp7α

149

149

Interacts with nsp9; viral RNA synthesis.

nsp7β

120

110

?

nsp8

45

45

?

883

1019

Down regulates Swine Leukocyte Antigen class I; PLP2 domain.

714

850

Potential innate immune antagonists; PLP2 domain.

645

646

RNA-dependent RNA polymerase; virus transcription and replication.

nsp10

442

441

RNA helicase/NTPase; putative zinc binding domain; role in subgenomic mRNA synthesis.

nsp11

224

223

Nidovirus uridylate-specific endoribonuclease (NendoU); IFN inhibition.

nsp12

152

153

Interacts with cellular chaperons

nsp2TF

ORF_TF

nsp2N nsp9

ORF1b

Structural proteins GP2

ORF2a

249

256

Small integral envelope protein, essential for virus infectivity, incorporated into virions as a multimeric complex, viral attachment protein, anti-apoptotic activity.

E

ORF2b

70

73

Minor un-glycosylated and myristoylated protein; possesses ion-channel like properties, essential for virus infectivity, and induces apoptosis.

GP3

ORF3

265

254

Minor glycoprotein, highly glycosylated with N-linked oligosaccharides, part of GP2/GP3/ GP4 heterotrimer.

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Table 7.1 Continued Number of amino acids Proteins

ORFs

Type I

Type 2

Function

GP4

ORF4

183

178

Minor glycoprotein highly glycosylated protein (four N-linked glycosylation sites), part of GP2/GP3/GP4 heterotrimer, viral attachment protein.

GP5

ORF5

201

200

Major structural protein, GP5/M heterodimer crucial for virus assembly, involve in the entry of virus into the host cells.

ORF5a protein

ORF5a

43

51

Involved in RNA-binding

M

ORF6

173

174

Major un-glycosylated structural protein, integral membrane protein, important for virus assembly and budding, part of GP5/M heterodimer.

N

ORF7

128

123

Genome packaging, localize in nucleus/ nucleolus, potential IFN antagonist.

Figure 7.2 PRRSV life cycle. The virus enters the cell via receptor-mediated endocytosis. Following uncoating, viral non-structural proteins (pp1a and pp1ab) are translated directly from the genome and assemble into replication and transcription complex (RTC) over double membrane vesicles. RTCs produce both full-length and subgenome-length negative strand RNA. These negative sense RNA intermediates become templates for full-length positive-sense genomic RNA and translation of non-structural proteins whereas the later sets of RNA become the template for subgenomic RNA that produces structural proteins. The newly synthesized genomic RNA is packaged into nucleocapsid and enclosed by an envelope along with other structural proteins in the Golgi. Finally, the virus is released by exocytosis.

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(RdRp; nsp9), the zinc-binding domain (ZBD; nsp10), the RNA helicase (nsp10), and the conserved nidovirus uridylate-specific endoribonuclease (NendoUorU; nsp11) (Ulferts and Ziebuhr, 2011). It has been demonstrated that the ORF1b (nsp9–12) is generated approximately once for every six translational events of the PRRSV genome, which is justifiable since these proteins are required only for the replication process (Kappes and Faaberg, 2015). The lack of a 3′ proof-reading ability of arteriviral RdRps makes them more prone to mutations in the genome. Consequently, an abnormally high evolution rate is estimated to be between 4.71 × 102 and 9.8 × 102/synonymous sites/year (Hanada et al., 2005). sgmRNA and heteroclite RNA Production of sg mRNA and heteroclite RNA are hallmarks of the PRRSV replication cycle. Structurally, sg mRNA has a ‘body’ that encodes viral genes derived from the 3′ end of the viral genome and a common ‘leader sequence’ derived from the 5′ UTR region (de Vries et al., 1990). The leader sequence contains the transcription regulatory sequences (TRS), a feature which is shared with coronaviruses. PRRSV produces six sg mRNAs (sg mRNA2–7) that encode for viral structural proteins. All sg mRNAs are genetically polycistronic (except sg mRNA7) but are functionally monocistronic. At the same time, sg mRNA2 and 5 are bicistronic and produce GP2 and E (sg mRNA2) and GP5 and 5a (sg mRNA5) proteins. These are synthesized discontinuously from the 3′-end and translocated and join the 5′ leader sequence (Lin et al., 2002; Meng et al., 1996). PRRSV produces a form of defective interfering RNA molecules, the heteroclite RNA. It is made up of short sequences of 3′-end of the genome and 5′-end of the ORF1a, excluding the sequence of 3′ and 5′ UTRs. The different sizes of heteroclite RNA are categorized as S1–9 based upon variable length of ORF1a, but maintain a common sequence (nt 191–476) (Yuan et al., 2000; Yuan et al., 2004). Heteroclite RNA molecules are co-packaged into the viral particles along with genomic RNA. A stretch of 35 nt on the heteroclite RNAs has the packaging signal that interacts with the nucleocapsid protein (Baig and Zakhartchouk, 2011). The production of these RNA molecules are a crucial part of the PRRSV life cycle, as they can be detected under all infection conditions and are thought to be responsible for the delayed and insufficient host immune response in PRRSV infection (Yuan et al., 2000). Nucleocapsid and RNA packaging Structurally, the PRRSV nucleocapsid is pleomorphic, mostly influenced by the shape of the outer envelope, and appears as a two layered shell with a thickness of about 10–11 nm. The core has a large central cavity. The N protein forms the nucleocapsid that packages the viral RNA. The C-terminus of the N protein contains a dimerization domain, causing the N protein to form homodimers. In the core, the N protein interacts with the viral RNA via the positively charged residues (35–51) in the N-terminal domain (Fig. 7.3) (Wootton et al., 2002; Yoo et al., 2003). Two layers of the N protein homodimer with RNA in the middle form a twisted chain that bundles into a loose spherical shape leaving a hole inside (Spilman et al., 2009).

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Figure 7.3 Nucleocapsid (N) protein and encapsidation model. (A) N protein comprises RNA binding domain (1 to 57 aa) and dimerization domain (57 to 123 aa). Part of the RNA binding domain is rich in basic amino acids (35 to 57aa) that interact with viral RNA genome. (B) 3D homodimeric structure of N protein. Each chain comprises three α-helixes, and two β-sheets arranged themselves as shown in the cartoon diagram (left). The RNA-binding amino acids (35 to 57) are highlighted by surface structure diagram (right). (C) Schematic model of organization of the viral nucleocapsid.

Cellular biology of the virus Cell tropism and receptors PRRSV has a restricted cell tropism, and it infects in vivo cells of the monocyte/macrophage lineage, such as differentiated macrophages in lungs, lymphoid tissues and placenta. In vitro, the virus can replicate in African green monkey kidney cell line MA-104 and its derivatives, such as MARC-145 and CL-2621 (Kim et al., 1993; Mengeling et al., 1995). Also, PRRSV can infect monocyte- or bone marrow-derived porcine dendritic cells, but not lung dendritic cells (Loving et al., 2007). Several factors have been described to be involved in the process of the virus infection of cells. First, PRRSV uses heparin sulfate on the cell surface for a low-affinity attachment (Delputte et al., 2002). Second, the viral GP5/M protein complex binds to the N-terminal portion of CD169 (sialoadhesin), which trigger receptor-mediated, claritin-dependent endocytosis (Delputte et al., 2007). Third, the viral particles are transported to early endosomes, and the viral genome is released into the cytoplasm with the help of acidification and a scavenger receptor CD163 (Welch and Calvert, 2010). Also, the protease cathepsin E plays a role in this process (Misinzo et al., 2008). The cysteine-rich domain 5 of CD163 interacts with GP2 and GP4 proteins of the virus (Das et al., 2010; Van Gorp et al., 2010). Several nonpermissive cell lines allow productive PRRSV infection upon expression of porcine CD163 (Welch and Calvert, 2010). In addition, pigs engineered by knockout of CD163, but not sialoadhesin, became resistant to PRRSV infection (Prather et al., 2013; Whitworth et al., 2016). These data indicate that CD163 receptor is a key player in the process of PRRSV infection in vivo. Among other possible receptors, simian vimentin and CD151 may play a role in PRRSV infection of MARC-145 cells (Kim et al., 2006; Shanmukhappa et al., 2007).

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Virus–host cell interactions With the limited resources of their own, viruses explore the host machinery for their propagation and sustenance, and manipulate different host signalling pathways in their favour. PRRSV also manipulates with multiple signalling pathways (Table 7.2) to promote viral infection of host cells and to combat anti-viral immune responses. The virus encodes proteins that interact with host cellular proteins to carry out this function. For instance, a portion of the N protein targets the nucleolus of infected cells by nuclear and nucleolar localization sequences (Rowland et al., 2003). Once in the nucleus, the PRRSV N protein interacts with host cell proteins fibrillarin and nucleolin, and may be involved in cell cycle regulation (Yoo et al., 2003). Moreover, N has been found to interact with a human I-mfa domain-containing protein (HIC) homologue, a zinc-binding transcriptional regulator (Song et al., 2009), suggesting that N plays a role in transcriptional regulation in PRRSV-infected cells. In addition, the N protein in association with nsp9 interacts with cellular DHX9 to regulate viral RNA synthesis (Liu et al., 2016). Recently, Jourdan et al. (2012) constructed an interactome map of the N protein using stable isotope labelling by amino acids in cell culture (SILAC)-based quantitative proteomics. This map includes numerous cellular proteins, mainly those involved in forming the translation initiation complex and splicing. In addition, Sagong and Lee (2010) characterized differential cellular protein expression in continuous PAM cells expressing the N protein. They found 15 proteins with significant alteration of expression, wh ich we re classified into the functions involved in a variety of cellular processes, such as cell division, metabolism, inflammation, stress response, transportation, and others. Similarly, Liu et al. (2015) identified PARP-1 as a cellular interactor for the N protein and showed that this interaction is critical for the virus replication. In a study by Wang et al. (2014a), the host cellular proteins that interact with nsp2 were immunoprecipitated with anti-Myc antibody from MARC-145 cells infected by a Table 7.2 Cell signalling pathways regulated by PRRSV Pathway

Reference

Extracellular signal-regulated kinase (ERK)

Lee and Lee (2010)

Phosphatidylnositol-3-kinase (PI3K)/Akt

Zhang and Wang (2010); Zhu et al. (2013)

c-Jun N-terminal kinase (JNK)

Yin et al. (2012)

Unfolded protein response (UPR); p53

Huo et al. (2013)

Retinoic acid-inducible gene I (RIG-I); activator protein-1 (AP-1)

Lou et al. (2008)

Toll-like receptors

Chaung et al. (2008)

Apoptosis

Lee and Kleiboeker (2007); Costers et al. (2008)

Autophagy

Liu et al. (2012)

Mammalian target of rapamycin (mTOR)

Pujhari et al. (2014)

Nuclear factor-κB (NF-κB)

Lee and Kleiboeker (2005); Fu et al. (2012)

Type I interferon response

Albina et al. (1998)

Tumour necrosis factor alpha (TNF-α)

López-Fuertes et al. (2000)

MicroRNA

Hicks et al. (2013)

|

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recombinant PRRSV with a Myc tag insertion in its nsp2-coding region. The 285 cellular proteins were identified by liquid chromatography-tandem mass spectrometry (LC-MS/ MS). Functional analyses of the interactome profile highlighted pathways associated with infectious disease, translation, the immune system, the nervous system, and signal transduction. Interactions of nsp2 with two cellular proteins (BCL2-associated athanogene 6 (BAG6) and apoptosis-inducing factor 1 (AIF1)) were validated by a co-immunoprecipitation assay. Interactome of nsp2 has also been characterized in two other studies (Song et al., 2016; Xiao et al., 2016). The data indicated that nsp2 interacted with a number of cellular proteins including 14-3-3, CD2AP and vimentin. Another study (Dong et al., 2014) demonstrated that nsp9 interacted with cellular retinoblastoma protein (pRb). The pRb level was confirmed to be down-regulated in PRRSV-infected MARC-145 cells, and nsp9 was shown to promote pRb degradation. The interactome of nsp12 has also been determined using quantitative proteomics coupled with an immune-precipitation strategy based on over expression of nsp12–EGFP fusion protein in human 293T cells (Dong et al., 2016). About 100 cellular proteins were identified, and interaction with HSP70 was verified. Taken together, all these data indicate existence of interactions between viral and cellular proteins, and these interactions are important for PRRSV replication and pathogenesis. Regulation of apoptosis Apoptosis is an important process by which virus-infected cells are eliminated from the host; therefore, viruses have evolved mechanisms to prevent or delay apoptosis. PRRSV stimulates anti-apoptotic pathways early in infection (Costers et al., 2008). In MARC-145 cells stably transfected with the GP2 encoding gene, staurosporine-induced apoptosis was inhibited suggesting a potential anti-apoptotic role of GP2 (Pujhari et al., 2014a). On the other hand, viruses might induce apoptosis to enhance spread of infection and avoid host immune responses. PRRSV also triggers apoptosis at the late stage of infection (Costers et al., 2008). PRRSV nsp4 and nsp10 were shown to be pro-apoptotic proteins, and nsp4 induced apoptosis through manipulation with the pro- and anti-apoptotic functions of the Bcl-2 family member proteins, whereas activation of caspase-8 and Bid are required for nsp10-induced apoptosis (Ma et al., 2013; Yuan et al., 2016). Also, E protein induced apoptosis by activation of caspase-3. The exact mechanism of E protein pro-apoptotic function needs to be further elucidated, although interactions between E and some mitochondrial proteins were demonstrated which led to decreasing the ATP level in the host cells (Pujhari and Zakhartchouk, 2016). Subversion of the type I interferon response The antiviral defence of the host is armed with a mechanism to detect and eliminate invading viruses. Double stranded RNA is sensed by pattern recognition receptors: toll-like receptor (TLR) or retinoic acid-inducible gene I (RIG-I). Activation of TLR3 leads to dimerization of receptors and recruitment of TIR-domain-containing adapter-inducing interferon-β (TRIF). This stimulation results in the assembly of signalling complexes and initiation of signalling cascades leading to the phosphorylation and activation of interferon regulatory transcription factor 3 (IRF3) and IRF7. On the other hand, RIG-I activates the IFN promoter stimulator-1 (IPS-1), and IPS-1 then induces signalling pathways resulting in the activation of IRF3/7, NF-kB and AP-1. Once activated, these transcription factors

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translocated to the nucleus, and together with CREB-binding protein (CBP), induce the transcription of type I IFN genes. Once IFNs are secreted extracellularly, they bind to the IFN receptor and the receptor associated JAK1 kinase is phosphorylated and activated to recruit signal transducer and activator of transcription 1 (STAT1) and STAT2. The phosphorylated STAT1 and STAT2 form IFN-stimulated gene factor 3 (ISGF3) complex along with IRF9. ISGF3 then undergoes nuclear translocation and induces the transcription of hundreds interferon stimulated genes (ISGs). PRRSV may inhibit IFN-α (Lee et al., 2004; Miller et al., 2004) and IFN-β (Overend et al., 2017) responses, and this inhibition is RIG-I mediated (Luo et al., 2008). Five PRRSV proteins were identified and characterized as IFN antagonists (Table 7.3). Cell membrane rearrangement PRRSV infection triggers the formation of double membrane vesicles (DMV) in which the viral replication and transcription complex (RTC) is thought to be located (Pedersen et al., 1999). The formation of DMV and convoluted membranes is a hallmark feature of cells infected with arterivirus (Knoops et al., 2012). Non-structural proteins with membrane spanning domains (nsp2, nsp3 and nsp5) are suggested to be responsible for induction of these membrane rearrangements (Fang and Snijder, 2010); however, the exact mechanism

Table 7.3 Type I IFN antagonistic proteins encoded by PRRSV and mechanisms of function Viral proteins

Mechanism of function

References

Nsp1α

Decreases IFN promoter activity

Chen et al. (2010)

Binds to and mediates CBP degradation

Kim et al. (2010)

Inhibits NF-κB signalling

Song et al. (2010)

Inhibits IFN promoter activity

Chen et al. (2010)

Inhibits IRF3 phosphorylation and nuclear translocation

Beura et al. (2010)

Inhibits host cellular mRNA export

Han et al. (2017)

Blocks STAT1/STAT2 translocation

Patel et al. (2010)

Induces degradation of karyopherin-α1 and blocks ISGF3 translocation

Wang et al. (2013)

Inhibits IRF3 phosphorylation and nuclear translocation

Li et al. (2010)

Suppress NF-κB activation by inhibition of IκB degradation

Sun et al. (2010)

Inhibits ISG15 production and ISG15-mediated ISGylation

Sun et al. (2012)

Inhibits IFN-β production

Chen et al. (2014)

Inhibits NF-κB signalling by cleaving NEMO

Huang et al. (2014)

Cleaves IPS-1

Huang et al. (2016)

Inhibits IFN promoter activity

Shi et al. (2011)

Suppress NF-κB and RIG-I signalling by cleaving IPS-1 mRNA

Sun et al. (2016)

Inhibits IRF3 phosphorylation and nuclear translocation

Sagong and Lee (2011)

Nsp1β

Nsp2

Nsp4

Nsp11

N

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of DMV formation has not been clarified yet. Although, since PRRSV induces autophagy to benefit its replication (Liu et al., 2012), the DMV may resemble autophagosomes. Pathogenesis The clinical features of PRRS include two sets of symptoms: reproductive failure in pregnant females and respiratory disease in young pigs (Table 7.4). Highly pathogenic PRRS virus (HP-PRRSV), that emerged in China in 2006, shows more severe clinical signs (Box 7.1). PRRSV has specific tropism for differentiated macrophages (CD169+ and CD163+), and can cause a persistent infection in pigs (Wills et al., 1997b). The virus can be detected in blood for long periods of time, but the length of viraemia is strongly dependent on the virus strain and the age of pig. It may extend to weeks in piglets (up to 90 days), whereas in adult pigs it may last for only a few days. After viraemia, PRRS virus can persist for several weeks, particularly in tonsils and other lymphoid tissues. Although most pigs clear PRRS virus within 120 days (Batista et al., 2002), some may remain persistently infected for several months, and virus may be isolated from tonsils up to 105 days post infection (Horter et al., 2002). Initially, virus replicates in alveolar macrophages and then spreads via blood to placenta and lymphoid organs, such as the tonsils and lymph nodes, and to other tissues but less consistently. The virus compromises macrophage function and provokes an inflammatory cell infiltration resulting in interstitial pneumonia which, in conjunction with enlarged lymph nodes, is the common lesion of respiratory PRRSV infection. Lesions can also be observed microscopically in the uterus of sows and in the testicles of boars. Clinical disease and lesions due to the PRRS virus infection in post-natal pigs occur by a variety of mechanisms including apoptosis of macrophages (Lee and Kleiboeker, 2007; Costers et al., 2008) and induction of inflammatory cytokines (Peng et al., 2009). In pregnant sows, PRRSV replicates and causes severe vasculitis in the endometrium in late gestation (Harding et al., 2017; Karniychuk and Nauwynck, 2013), and rapidly crosses the maternal uterine epithelium and fetal trophoblast before infecting tissues (Suleman et al., 2018). Although compromised fetuses cluster in litters, the exact mechanisms of fetal Table 7.4 Clinical signs of PRRS Pigs

Clinical signs

Sows

Depression and lethargy; mild fever; coughing and respiratory signs, cyanosis of the ears and vulva; abortion at late stage of gestation; birth of small, weak, stillborn and mummified piglets; difficulty conceiving and delayed returns to estrus;

Boars

Depression and lethargy; mild fever; weight loss; lack of libido and decreased semen quality;

Pre-weaning piglets

Born weak; increased mortality in the first week of life; swollen conjunctiva and eyelids; apathy, emaciation from lack of food intake and splay leg; increased susceptibility to respiratory and systemic coinfections;

Wean-to-finish pigs

Lethargy and anorexia; dyspnoea and interstitial pneumonia; reductions in average daily gain and feed conversion; increased mortality; increase in secondary infections (respiratory and systemic)

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Box 7.1  Clinical signs of highly pathogenic PRRS virus in pigs High fever Depression, anorexia and lethargy Respiratory disorders, cough and dyspnoea Conjunctivitis Hyperaemia or cyanosis of extremities Haemorrhagic lesions: lung, liver, kidney and lymph nodes Increased in secondary bacterial infections Abortions (40%) High mortality (20–100%)

death are not fully understood, but may include apoptosis of fetal implantation sites, placental separation, pro-inflammatory cytokine expression, fetal hypoxia, and other direct effects of the virus on fetal tissues (Karniychuk et al., 2011; Ladinig et al., 2015; Novakovic et al., 2016, 2017; Wilkinson et al., 2016; Harding et al., 2017). Recent evidence also suggests that variability in fetal susceptibility may be related to polymorphisms in a number of regions in the fetal host genome (Yang et al., 2016). Because PRRSV can affect cells of the immune system, this may allow secondary pathogens to cause more serious disease. PRRSV is the most common infectious agent of porcine respiratory disease complex (PRDC); a multifactorial respiratory disorder of growing pigs characterized by anorexia, slow growth, fever, cough, and dyspnoea. Clinical disease was exacerbated in pigs co-infected with PRRSV and Mycoplasma hyopneumoniae (Thacker et al., 1999), and PRRSV and Bordetella bronchiseptica (Brockmeier et al., 2000). In addition, pigs infected with both PRRSV and porcine circovirus type 2 (PCV-2) showed more severe clinical symptoms and lung lesions than those associated with infection by either agent alone (Allan et al., 2000). Also, PRRSV infection exacerbates the inflammatory response to porcine respiratory coronavirus in pigs (Renukaradhya et al., 2010). Immune dysregulation by PRRSV PRRSV has a negative impact on the innate immune responses. Extremely low levels of interferon alpha (IFN-α) were detected in serum and lung secretion of pigs infected with PRRSV strains (Albina et al., 1998b). Also, PRRSV suppressed tumour necrosis factor-alpha production in infected macrophages (López-Fuertes et al., 2000), although this production varies significantly between the viral strains (Gimeno et al., 2011). Certain PRRSV strains were also able to induce regulatory cytokines, such as interleukin 10 and transforming growth factor-β (Chung and Chae, 2003; Silva-Campa et al., 2012). Infection of natural killer (NK) cells with certain PRRSV strains induced significant suppression of their cytotoxic activity (Renukaradhya et al., 2010). It has been demonstrated that PRRSV can infect monocyte- or bone marrow-derived porcine dendritic cells (Loving et al., 2007), and this infection led to reduction of MHC class I and II molecules expression and cell death via both apoptosis and necrosis (Rodríguez-Gómez et al., 2013). As a consequence of these events, humoral and cell-mediated immune responses to PRRSV are adversely affected and delayed. Antibodies against PRRSV can be detected very rapidly, as soon as 5 days post infection, but these early antibodies are mainly directed to the N protein and do not possess the

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virus-neutralizing activity. Virus-neutralizing antibodies (VN) can be detected around one month or even later. Nevertheless, VN titres against PRRS virus are usually low in comparison with those induced by other viruses (Loemba et al., 1996). PRRSV infection induces the development of a slow, erratic and low frequency of interferon gamma (IFN-γ) secreting cells. IFN-γ-secreting cells were mainly CD4+ CD8+ cells, with a few CD4–/CD8αβ+ cytotoxic T cells (Meier et al., 2003). The frequency of Foxp3+ T-regulatory cells, which are immunosuppressive, was increased (Silva-Campa et al., 2012). Diagnostics Within and between herds, the severity of PRRS virus infection can vary widely from lack of clinical signs to devastating outbreaks; therefore, a PRRS diagnosis based solely on clinical signs can be extremely complicated or lead to error. This is particularly true in PRRSV-positive farms that are re-infected with a heterologous strain of PRRSV. Herd-level diagnosis involves investigation of many components such as production records, clinical history, clinical signs, serology, pathology and virus detection. Changes in a number of reproductive parameters identified in standard herd production records, such as increases in late term abortions, early farrowings, and birth of mixed litters containing weak-living, stillborn and mummified fetuses are indicative of reproductive PRRS. Likewise, increased pre-weaning mortality, non-antibiotic neonatal diarrhoea, and respiratory problems, particularly dyspnoea, in lactating piglets may suggest the occurrence of in utero PRRSV transmission. In endemically infected herds, pre-existing immunity resulting from natural infection, vaccination, or both, generally prevents transplacental infection, and passive immunity generally protects pigs until 6–8 weeks of age. PRRSV exposure after the decay of maternally derived antibodies results in infection that can be asymptomatic or cause respiratory disease with severity dependent on the PRRSV strain and presence of other respiratory pathogens. Laboratory diagnosis is absolutely necessary to confirm PRRS infection. Numerous methods are widely used to detect or isolate virus in serum or tissues, including immunohistochemistry (IHC), immunofluorescent assay (IFA), reverse transcription polymerase chain reaction (RT-PCR; conventional and quantitative). Virus can be isolated in both porcine alveolar macrophages and sublines of the African monkey kidney cell line (CL-2621 and MARC-145). Since cytopathic effects, which occur in one to five days, are not always evident, virus isolation usually needs to be confirmed by RT-PCR, IHC or IFA. Sequence analysis of the ORF5 region of PRRSV genome is routinely used for strain identification and epidemiologic investigation. To detect virus-specific antibodies appropriate tests include enzyme-linked immunosorbent assay (ELISA), immunoperoxidase monolayer assay (IPMA), immunofluorescent antibody assay (IFA) and virus/serum neutralization test (VNT or SNT). ELISA is the most commonly used because it is easy to perform, rapid and cheaper compared to other assays. Unfortunately, none of the serological tests can distinguish between the wild-type virus and modified live virus vaccines. Recently, oral fluid samples have been evaluated as an alternative to serology (Langenhorst et al., 2012).

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Epidemiology The origin of the virus is not known, but some have proposed that it came from a wild pig reservoir (Plagemann, 2003). Nowadays, PRRS is prevalent in most countries and exists in epidemic and endemic forms. PRRS is considered as one of the most costly diseases to the worldwide pig industry, and it is estimated to cause $664 million economic losses in the USA annually (Holtkamp et al., 2013). Additionally, highly pathogenic strains emerged in China in 2006, affected over 2 million pigs within a few months, with mortality rates of 20–100% in infected populations. As previously mentioned, there are two distinct species of PRRSV (Adams et al., 2016): PRRSV-1 (European, prototype Lelystad virus) and PRRSV-2 (North- American, prototype VR-2332). High genetic diversity is a hallmark of PRRSV. Based on ORF-5 sequences, several subtypes or clades of the virus have been identified in both species. There are at least four different subtypes of PRRSV-1, whereas PRRSV-2 consists of nine lineages of isolates (Shi et al., 2010). Presently, both PRRSV species are disseminated worldwide. One or both species can be found in almost all pig-producing countries, with the exception of Australia, New Zealand, Scandinavia, Switzerland and some South American countries. Virulence varies within and between both species; extremely virulent strains being the highly pathogenic PRRSV-2 that emerged in China and rapidly spread throughout other Asian counties, and the highly pathogenic PRRSV-1 that was isolated from a Belarusian farm (Karniychuk et al., 2010). Transmission between herds The most commonly accepted way for disease to be transmitted between herds is by use of contaminated semen or by the introduction of infected animals. A reported breakdown (Le Potier et al., 1997) of the sources of herds’ infection is presented in Fig. 7.4. Transmission by aerosol is possible and virus captured by air filtration about 9 km from an experimentally infected farm was shown to be viable indicating contaminated aerosols may play an important role in regional transmission (Otake et al., 2010). Aerosol transmission, however, appears to be strain­specific (Cho et al., 2007) and depends on meteorological conditions (Hermann et al., 2007). Transmission of PRRSV has been demonstrated experimentally via houseflies and mosquitoes. The virus can survive for a short period on the exterior surface of the insects whereas the gastrointestinal tract being the main site of retention of the virus (Rochon et al., 2015). Rodents (mice and rats) are not a reservoir for the virus (Hooper et al., 1994).

Figure 7.4 Estimated sources of PRRSV infection for herds in France (Le Potier et al., 1997).

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Mallard ducks are susceptible to PRRSV infection and can excrete the virus in their faeces for up to 39 days post experimental infection (Zimmerman et al., 1997). Transmission within herds Infected pigs can shed virus from multiple routes and for extended periods of time, mainly in nasal secretions, saliva and semen, less frequently in mammary secretions, and rarely in urine and faeces (Wills et al., 1997a). The amount of virus shed and the duration of the shedding vary significantly among strains. Overall, prolonged viraemia and persistent infection increase the possibility of PRRSV transmission. Congenitally infected piglets are an important reservoir of PRRSV that can be transmitted to siblings or other susceptible cohort piglets, and therefore must be prevented in developing an effective PRRS control strategy. PRRSV vaccines The first commercial PRRSV vaccine was released in the USA in 1994. However, in spite of the use of this and subsequent vaccines, PRRSV continues to cause substantial economic losses in swine industry. Obstacles for the development of effective vaccines against PRRSV include high genetic diversity of the virus, modulation of the host immune system, interfering with neutralizing antibodies by decoy epitopes and glycan shielding of epitopes, incomplete understanding of the correlates of protection, and lack of reliable parameters to predict vaccine protective efficacy (Kimman et al., 2009; Murtaugh and Genzow, 2011; Vu et al., 2016). Modified live PRRSV vaccines Currently, at least 10 PRRSV modified live virus (MLV) vaccines are licensed for use in countries around the world. The development of MLV vaccines involves virus attenuation by multiple passages through cell culture. During this process, the virus gradually accumulates mutations that allow it to grow better in its new host cell, simultaneously losing virulence in pigs. Almost all PRRSV MLV vaccines were developed by passaging the virus strains on African green monkey kidney cells. There is only one exception: Fostera PRRS MLV vaccine (Zoetis Animal Health, NJ, USA) that was developed by attenuating PRRSV-2 by passage on CD163-expressing swine and hamster cell lines. Attenuated PRRSV vaccine strains can infect lung, lymphoid tissues and placenta, and it is transmissible to contact pigs and through semen. Therefore, MLV vaccines are not approved for use in PRRSV virusnegative herds, pregnant females, and breeding age boars. MLV vaccinated pigs elicit neutralizing antibody and cell-mediated immune responses, although these responses are delayed and relatively weak (Meier et al., 2003; Zuckermann et al., 2007). The duration of immunity is at least four months, roughly equivalent to the finishing period or gestation. MLV vaccines do not provide full protection against PRRS virus infection, but they can reduce the impact of the disease. In growing pigs, vaccination with MLV reduced viraemia, respiratory signs, macroscopic and microscopic lung lesions (Park et al., 2014). Also, vaccination can reduce wild type PRRS virus shedding from an infected population of growing pigs (Linhares et al., 2012) and protect piglets from lethal challenge with HP-PRRSV (Tian et al., 2009). In PPRSV-infected sows, MLV vaccination helps to reduce abortions, and increase farrowing rate and number of weaning pigs (Alexopoulos et al., 2005).

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There is a general assumption that MLV vaccines confer complete protection against homologous PRRSV strain, whereas heterologous protection is variable (Murtaugh and Genzow, 2011). However, protection afforded by a given vaccine or a given field strain against another strain cannot be predicted simply by the genetic similarity (Prieto et al., 2008). There are safety concerns of PRRSV MLV vaccines because of the risk of the vaccine reverting to virulence under field conditions. The reversion of a commercial MLV to a pathogenic phenotype and the possibility of recombination with field isolates have been demonstrated (Bøtner et al., 1997; B. Li et al., 2009a). Therefore, there is a clear need for more safe and effective MLV vaccines. Strategies to improve PRRSV MLV vaccines Several strategies have been employed to broaden the cross-protective efficacy of PRRSV MLV vaccines. All these studies were possible after the generation of full-length infectious clones of PRRSV (Nielsen et al., 2003; Lee et al., 2005). Wang et al. (2008) constructed chimeric recombinant viruses in which the 5′ UTR/ORF1 of one strain genome was linked to ORF2–7/3′ UTR from the other strain genome. Using similar approach, a PRRSV chimeric virus was constructed in which ORFs 3–4 and 5–6 were replaced with two Korean field isolates. The chimeric virus conferred a broader range of protection against challenges with two heterologous viruses (Shabir et al., 2016). Another approach to broaden the cross-protective efficacy of the MLV vaccine is DNA shuffling of the viral genes encoding structural capsid proteins. Molecular breeding through DNA shuffling rapidly generates recombinant sequenced in vitro. To this end, ORF3, ORF4, ORF5 and ORF6 gene fragments from each of the six parental PRRSV strains were digested by DNase I and reassembled by PCR. In one of the studies, the ectodomain sequences of these genes in different combinations were incorporated into an infectious clone of PRRSV MLV. Five viable progeny chimeric viruses were rescued, and one of the viruses conferred partial cross-protection against challenge with two heterologous PRRSV strains (Tian et al., 2015). In a subsequent study, the full-length individually shuffled structural genes were assembled into the backbone of PRRSV strain VR-2385, and recombinant virus was shown to induce cross-protection in pigs challenged with two heterologous strains (Tian et al., 2017). In another study (Vu et al., 2015), the authors used a centralized immunogen approach to expand the antigen coverage of a PRRSV MLV vaccine. A consensus PRRSV genome was designed based on alignment of 59 non-redundant full-length genome sequences of PRRSV-2 and selecting the most common nucleotide found at each position of the alignment. This consensus PRRSV genome was chemically synthesized, and a full-length infectious cDNA clone was constructed. When inoculated into pigs, the recombinant virus with consensus genome sequence conferred significantly broader levels of protection against challenge with a heterologous PRRSV strain. Currently, it is impossible to differentiate between pigs infected with field strain or pigs vaccinated with MLV vaccine. This problem can be solved by developing serological marker vaccines, which there are two types. First, DIVA (Differentiating Infected From Vaccinated Animals) vaccines involve the elimination of one or more antigenic epitopes from the vaccine strain that remain present in field strains. Second, compliance marker vaccines, involve the insertion of antigenic epitopes into the vaccine strain that makes it possible to detect this

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strain among all other field strains. Both approaches have been used in PRRSV MLV vaccine development by making appropriate insertions or deletions in PRRSV full-length infectious clones (Fang et al., 2008; Lin et al., 2012); however, there are presently no commercially available DIVA or compliance marker PRRSV MLV vaccines licensed for use in commercial farms. Killed PRRSV vaccines PRRS killed virus (KV) vaccines are licensed for use in many countries around the world excluding North America. Commercial PRRSV KV vaccines are considered less efficacious than MLV vaccines in inducting immune responses and protection (Zuckermann et al., 2007). However, administration of a KV PRRSV-1 vaccine to gilts and sows in a seropositive herd for period of 18 months resulted in significant improvement in reproductive performance and an increased percentage of weaned pigs (Papatsiros et al., 2006). In another study (Scortti et al., 2007), breeding gilts were vaccinated with KV vaccine and challenged with a heterologous PRRSV-1 strain. The vaccine failed to prevent the clinical signs of disease, viraemia and transplacental infection of piglets; however, the pre-weaning mortality was significantly reduced (Scortti et al., 2007). The major advantage of a KV PRRSV vaccine is its safety. To date, there have been no reports of negative impacts of PRRSV KV vaccines on pig health. Strategies to improve PRRSV KV vaccines Proper methods of virus inactivation and suitable adjuvants in are critical for improving the efficacy of PRRSV KV vaccines. In vitro experiments with porcine macrophages demonstrated that binary ethylenimine (BEI) and UV-radiation are ideal methods to inactivate PRRSV particles without losing their capacity to bind, internalize and disassemble (Delrue et al., 2009). Vaccination of pigs with PRRSV-1 inactivated with UV or BEI and formulated with oil-based adjuvants elicited increased VN titres and reduced viraemia following homologous virus challenge (Vanhee et al., 2009). In another study, vaccination of pregnant sows with BEI-killed PRRSV also elicited increased VN titres and reduced viraemia following homologous virus challenge. Although the vaccine did not prevent transplacental infection of fetuses, it did reduce the number of fetuses with severe microscopic lesions at the fetal implantation sites (Karniychuk et al., 2012). It has been reported that glycan shielding may interfere with the neutralizing antibody responses against PRRSV (Ansari et al., 2006). Using reverse genetics, researchers produced recombinant virus containing double-amino acid substitutions at the potential N-glycosylation sites of the GP5 protein. The PRRSV-2 was BEI-inactivated and formulated with the Montanide adjuvant to produce a KV vaccine. The efficacy of the inactivated mutant virus vaccine in pigs was compared with that of the inactivated wild-type virus. Overall, KV vaccine based on mutant virus increased the VN antibody response, reduced viraemia and lung lesions after homologous virus challenge (Lee et al., 2014). Mucosal vaccination offers attractive advantages to parenteral vaccination, such as local immune responses at the airway surface and in the lungs. Dwivedi et al. (2012) entrapped PLGA [poly (lactide-co-glycolides)] nanoparticles with UV-killed PRRSV-2 and administered intranasally to pigs. Vaccinated pigs showed increased virus-specific IgA levels in the lungs, increased VN titres, and high levels of IFN-γ production. After heterologous PRRSV

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strain challenge, pigs showed a significant reduction in lung pathology and virus load. In a continuation study, PLGA nanoparticle-based UV-killed PRRSV vaccine was co-administered twice intranasally with a potent adjuvant: Mycobacterium tuberculosis whole-cell lysate. Vaccinated pigs were challenged with heterologous PRRSV-2 strain, and complete clearance of challenged virus was reported with 3-fold reduction in viral RNA in the blood. Additionally, vaccinated pigs showed enhanced PRRSV-specific antibody immune responses including VN antibody titres, a balanced Th1/Th2 type response, decreased expression of immunosuppressive cytokines, increase frequency of IFN-γ-expressing lymphocyte subsets, and expanded population of antigen-presenting cells (Binjawadagi et al., 2014). Experimental PRRSV vaccines Numerous efforts have been made to develop a novel PRRSV vaccine with improved safety and protection efficacy. This includes construction of alternative vaccines such as DNA vaccines, subunit vaccines, synthetic peptide vaccine (Mokhtar et al., 2017) and virus-vectored vaccines. Non-comprehensive lists of these vaccines and brief summaries of research findings are presented in Tables 7.5–7.7. However, there is little evidence that these vaccine candidates provide substantial protection against PRRSV challenge outside small experimental studies. Moreover, in the majority of the experimental studies only homologous virus was in use in the challenge experiments. One exception is the PRRSFREE subunit vaccine from Reber Genetics Co. Ltd. (Taipei, Taiwan). This vaccine was tested in an infected herd of 300 sows in a farrow-to-finish farm in Taiwan (Yang et al., 2013). In 2012, this new recombinant PRRSV vaccine was introduced into the Taiwanese market. A follow up study demonstrated that this vaccine provided protection against respiratory disease from heterologous PRRSV-1 and PRRSV-2 challenge in growing pigs ( Jeong et al., 2017). Table 7.5 DNA vaccines Antigen

Model

GP5

Mice, pigs

G5, N plus IL-2 and IFN-γ Pigs

VN antibody

CMI

Protection

Reference

+

+

+

Pirzadeh and Dea (1998)

nt

+

+

Xue et al. (2004)

GP5 plus PADRE epitope

Mice, pigs

+

+

nt

B. Li et al. (2009b)

GP5 and M

Pigs

+

+

+

Chia et al. (2010)

GP5 plus IL-18

Pigs

+

+

nt

Zhang et al. (2012)

GP5, GP3 plus IFN-α/γ

Pigs

+

+

+

Du (2012)

GP5 plus IL-15

Mice

+

+

nt

Li et al. (2012)

GP5 plus ubiquitin

Pigs

+

+

+

Hou (2008)

PRRSV cDNA with deletion in ORF2

Pigs

–

+

+

Pujhari et al. (2013) Zhang et al. (2012)

GP5, M plus IL-18

Pigs

–

+

nt

GP5, M, N

Pigs

–

+

– enhanced Diaz et al. (2013) fever

+, detected; –, not detected; nt, not tested; VN, virus neutralizing; CMI, cell-mediated immunity.

168  | Zakhartchouk et al.

Table 7.6 Vectored vaccines Vector

Antigen

Model

VN antibody

CMI

Protection

Reference

Adenovirus

GP5, M

Mice

+

+

nt

Jiang et al. (2006)

Adenovirus

GP5, GP3 plus CD40L

Pigs

+

+

+

Cao et al. (2010)

Adenovirus

GP5, GP3 plus GM-CSF

Pigs

+

+

+

Wang et al. (2009)

Adenovirus

GP5, GP3, GP4

Mice

+

+

nt

Jiang et al. (2008)

Adenovirus

GP5, GP3 plus HSP70

Pigs

+

+

+

J. Li et al. (2009)

Baculovirus

GP5, M

Mice

+

+

nt

Wang et al. (2007)

Fowlpox virus

GP5, GP3 plus IL-18

Pigs

+

+

+

Shen et al. (2007)

Vaccinia (rMVA)

GP5, M

Mice

+

+

nt

Zheng et al. (2007)

Pseudorabies virus

GP5

Pigs

+

nt

+

Qiu et al. (2005)

M. bovis (BCG)

Truncated GP5; M

Pigs

+

–

+

Bastos et al. (2004)

Circovirus type 1

Epitopes of GP5, Pigs GP3, and GP2

+

nt

nt

Piñeyro et al. (2016)

Coronavirus (TGEV)

GP5, M

+

nt

+

Cruz et al. (2010)

Pigs

+, detected; –, not detected; nt, not tested; VN, virus neutralizing; CMI, cell-mediated immunity.

Table 7.7 Subunit vaccines Antigen

Production system

Model

VN antibody

CMI

Protection

Reference

GP5

Sf9 cells infected Mice with baculovirus

+

+

nt

Wang et al. (2012)

GP5, GP4, GP3, GP2a and M

Sf9 cells infected Pigs with baculovirus

nt

+

+

Bijawadagi et al. (2016)

Multiple B-cell epitopes of GP5, GP2, GP3, N and nsp2

E. coli

+

+

nt

Chen et al. (2012)

Mice, pigs

M

corn

Mice

+

+

nt

Hu et al. (2012)

*Pseudomonas

E. coli

Pigs

+

+

+

Yang et al. (2013) Jeong et al. (2017)

exotoxin carrier with epitopes of ORF1b, N, M and GP5

+, detected; –, not detected; nt, not tested; VN, virus neutralizing; CMI, cell-mediated immunity. *Commercial name of this vaccine is PRRSFREE.

Porcine Reproductive and Respiratory Syndrome Virus |  169

Conclusions Since the discovery of PRRSV in 1991, much has been learned about this virus. Thanks to the efforts of many scientists around the world, we have gained a better understanding of the virus molecular biology, pathophysiology, epidemiology, and virus–host cell interactions. The complete genomes of different PRRSV strains have been sequenced, and a reverse genetics system for PRRSV was established by construction of full-length infectious cDNA clones. New, more effective, diagnostic procedures have been developed. Based on improved knowledge of PRRSV, scientists and veterinarians developed the concept of ‘herd closure’ as a tool to eliminate PRRSV from herds. This approach is being used in ‘regional control and elimination programmes’ throughout the USA and Canada in an effort to eliminate PRRSV from less hog-dense swine population areas. A combination of herd closure and depopulation/repopulation methods helped to eliminate PRRSV in Chile and Sweden. The first commercially available PPRSV vaccine was licensed in the USA in 1994, and various anti-PRRSV vaccines are now widely used throughout swine producing areas in the world; however, safer and more effective PRRSV vaccines are needed. New knowledge about genetic diversity of the virus, viral modulation of the host immune system, and understanding of the correlates of protection will help to develop such a vaccine offering broad, heterologous protection. References

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Swine Vesicular Disease Virus Estela Escribano-Romero1, Miguel A. Martín-Acebes1, Angela Vázquez-Calvo1, Emiliana Brocchi2, Giulia Pezzoni2, Francisco Sobrino3* and Belén Borrego4

8

1Departmento de Biotecnología, Instituto Nacional de Investigación y Tecnología Agraria y

Alimentaria (INIA), Madrid, Spain.

2Istituto Zooprofilattico Sperimentale della Lombardia e dell’Emilia Romagna (IZSLER), Brescia,

Italy.

3Centro de Biología Molecular ‘Severo Ochoa’ (CSIC-UAM), Madrid, Spain. 4Centro de Investigación en Sanidad Animal (CISA-INIA), Madrid, Spain.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190913.08

Abstract Swine vesicular disease (SVD) virus (SVDV) belongs to the Enterovirus genus within the Picornaviridae family. This virus is genetically and antigenically highly related to the human coxsackie virus B5 (CVB5). Indeed, it has been shown that SVDV is a subspecies of CVB5 that arose as a result of an adaptation to swine. SVDV causes a vesicular disease that affects pigs, resulting in lesions and clinical signs similar to those of foot-and-mouth disease (FMD). In this chapter we have addressed different aspects relevant to understand the infectious cycle of SVDV, including its genome organization and the control of gene expression, the proteins encoded by the SVDV RNA and their known functions, as well as the role they play on cell entry and virus replication and pathogenesis. In addition, the characteristics of the virus particles and the adaptive response elicited by this virus, as well as current strategies for SVDV control by vaccination and other antiviral strategies are discussed. The clinical signs and lesions characteristic of SVD are also addressed, as well as the current approaches to its diagnosis, with special emphasis in its differentiation from FMD. Finally, an overview regarding SVDV control and epidemiology is also presented. Swine vesicular disease virus (SVDV) Swine vesicular disease is a usually subclinical or mild vesicular disease of pigs that closely resembles foot-and-mouth disease (FMD). It was first seen in Italy in 1966 and since then it has only been reported in Europe and a few Asian countries (Taiwan, Honk Kong, Macau and Japan). The aetiological agent of this disease, SVD virus (SVDV), is a positive-sense, single-stranded RNA virus that belongs to the Enterovirus genus in the Picornaviridae family (reviewed in Dekker, 2000).

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SVDV is considered to have recently arisen from the human enterovirus CVB5 as a result of an adaptation to swine. By means of phylogenetic studies, the divergence from a common ancestor has been estimated between 1945 and 1965 (Zhang et al., 1999). Clinical differentiation between SVD and FMD is difficult because of the similarity of their lesions and clinical signs. This difficulty, despite the mild course of the disease, has prompted the study of SVDV biology with special emphasis in the development of laboratory strategies to improve its diagnosis and characterization; for current information on SVD see (http://www.oie.int/fileadmin/Home/eng/Animal_Health_in_the_World/ docs/pdf/Disease_cards/SWINE_VESICULAR_DISEASE.pdf; and http://www.oie. int/eng/A_FMD2012/docs/2.01.05 FMD.pdf). Molecular biology of SVDV Genome structure The SVDV genome is composed of a single-stranded RNA molecule of positive polarity that is polyadenylated at its 3′ end and shows its 5′ end covalently linked to a copy of the viral protein 3B (VPg) (Fig. 8.1). This RNA molecule, of about 7400 nucleotides in length, excluding the poly(A), encodes a single polyprotein that is flanked by two highly structured untranslated regions (UTRs) located at the 5′ and 3′ ends of the genome. This genome organization is similar to that of the prototypic enterovirus, poliovirus (PV) (Inoue et al., 1989; Zhang et al., 1993). Both 5′ and 3′ ends of the RNA molecule are key players for viral replication and translation. Accordingly, the 5′ and 3′ non-coding regions are conserved between SVDV and coxsackieviruses, although there are several divergent nucleotide stretches (Inoue et al., 1989). It has been proposed that differences at the RNA 5′ end could be in part responsible for alterations in pathogenicity between viral strains (Seechurn et al., 1990), which have been associated with modifications in the efficiency with which the RNA is translated (Seechurn et al., 1990). In fact, translation of the unique open reading frame (ORF) is driven by an initiation mechanism independent of CAP, mediated by the internal ribosome entry site (IRES) element located within the 5′ UTR (Chen et al., 1993). Although the sequence of this IRES is relatively conserved among SVDV isolates, which indeed may be useful for diagnostics purposes, sequencing of different isolates revealed considerable nucleotide variability in the spacer region located between a cryptic AUG present at 39 nucleotides from the 3′ end of the IRES and the functional initiation codon of the polyprotein (Shaw et al., 2005). Moreover, some isolates, mostly from European origin, carry block deletions in this region, which could be related to their reduced growth ability in vitro (Shaw et al., 2005). All these findings point to the 5′ UTR as a key determinant on SVDV pathogenicity. SVDV proteins The SVDV RNA encodes a polyprotein of 2185 amino acids (Inoue et al., 1989; VázquezCalvo et al., 2016b) that is proteolytically processed in the cytoplasm of the infected cell (Fig. 8.1), rendering the different viral proteins (Inoue et al., 1989; Zhang et al., 1993; Escribano-Romero et al., 2000). The viral ORF can be divided in three regions: P1, P2 and P3. The P1 region encodes the four structural or capsid proteins 1A (VP4), 1B (VP2), 1C (VP3) and 1D (VP1). P2 and P3 regions encode the seven non-structural proteins (NSPs) (2Apro, 2B, 2C, 3A, 3B, 3Cpro,

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Figure 8.1  Genomic organization and polyprotein processing in SVDV. The scheme represents the single ORF (boxes) of SVDV flanked by the two UTRs located at 5′ and 3′ ends (lines). The lower part of the figure represents the polyprotein processing to render the mature proteins (see text for details).

and 3Dpol) involved in replication, translation and processing of the viral RNA (Zhang et al., 1993; Greninger, 2015). Regarding the necessary polyprotein processing to render the mature proteins, the cleavage between P1 and P2 regions is produced by the protease 2A (2Apro). The protease 3Cpro is in charge on the rest of polyprotein cleavages to yield mature proteins, except for the cleavage of the precursor VP0 to produce mature VP4 and VP2, which is essential for SVDV infectivity (Rebel et al., 2003). As proposed for other picornaviruses, VP0 cleavage is expected to be autocatalytic and dependent on RNA encapsidation (Hindiyeh et al., 1999). Capsid structure, structural proteins The capsid of SVDV virion is composed of 60 protomers of each of the four capsid proteins VP1–VP4, after the processing of the P1 precursor polypeptide (Fig. 8.2). The protomers self-assemble to form a non-enveloped icosahedral shell of diameter 25–30 µm and symmetry T = 1, which encloses the viral genome (Escribano-Romero et al., 2000). SVDV particles show a buoyant density of 1.30–1.34 g/ml in caesium chloride gradients, are resistant to ether and are stable throughout a wide range of pH (3–10) (Moore, 1977), which seems an adaptation to survive passage through the acidic conditions of the stomach. In a first step, the precursor P1 is processed by a viral protease to give three polypeptides, VP0, VP3 and VP1 that are assembled giving rise to the protomer (Fig. 8.2A and B). Five of these subunits or protomers are associated to form pentameric intermediates (Fig. 8.2C and D) and 12 of them self-assemble to form the capsid (Fig. 8.2E and F). The 12 pentamers may form an ‘empty capsid,’ or provirion, which lacks RNA and is considered a late intermediate in the morphogenesis of these viruses. Subsequently, the RNA molecule is introduced by a mechanism that is not well characterized. The last step of this assembly, once the RNA is already encapsidated, consists of the proteolysis, which – as mentioned above – is considered autocatalytic, of the precursor VP0 leading to VP2 and VP4. This proteolysis is

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Figure 8.2  Organization of SVDV capsid. (A and B) Outside (A) and inside (B) views of a protomeric subunit of SVDV capsid. Secondary structure is indicated by ribbon diagrams. (C and D) Outside (C) and inside (D) views of a pentameric subunit of SVDV capsid. Amino acid side chains have been omitted for clarity. (E and F) Different views of the SVDV capsid in diverse orientations. The atomic coordinates of SVDV capsid (Protein Data Bank code 1MQT) were used (Verdaguer et al., 2003). Structures were visualized using PyMol Molecular Graphics System version 1.5.0.4 (Schrödinger, LLC). VP1 in blue, VP2 in green, VP3 in red, and VP4 in yellow.

necessary to produce infective virus (Rebel et al., 2003). It has been reported the in vitro construction of recombinant SDV virus-like particles (VLPs) with uncleaved VP0 (Ko et al., 2005; Xu et al., 2017) for their use as antigen to detect antibodies against SVDV. In the mature virion VP1, VP2 and VP3 proteins (33, 32 and 29 kDa, respectively) are exposed at the viral surface forming a compact protein shell, whereas VP4 (about 7.5 kDa) lies across the inner surface of the capsid in contact with the RNA and thus not accessible from the outer shell surface. VP1–VP3 possess a tertiary structure consisting of a central hydrophobic nucleus of eight antiparallel β-sheets structures or β-barrel connected by variable extension loops, many of which are exposed to the surface of the virion. The C-termini of the three proteins are also oriented towards the surface, while their N-termini face its interior. Finally, there are also two internal α-helices, named αA and αB (Escribano-Romero et al., 2000). The atomic structure of SVDV from two isolates, UK/27/72 (Fry et al., 2003) and a more recent one, SPA/2/93 (Verdaguer et al., 2003) has been determined, confirming that it is more similar to CBV3 than to CAV9 and allowing to map the surface changes occurring during the adaptation of coxsackievirus B5 (CVB5) to infect pigs.

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Capsid proteins VP1 This protein comprises 283 residues, is the most variable of the four viral structural proteins and makes up most of the outer surface area of the capsid. It associates to form each of the 5-fold vertices of the icosahedral particle protruding from the rest of the capsid. There is a hydrophobic pocket inside the β-barrel of VP1 that in the rest of picornavirus contains lipids derived from cells and which is known as ‘pocket factor’. Its role is stabilizing the virus particle at the extracellular environment. In fact, removal of the pocket factor let the virus structure to ‘breathe’, probably allowing conformational changes necessary for uncoating (Giranda et al., 1992). In SVDV, there is still no chemical identification for this molecule as the size and shape of the density in its 3D structure is different from that observed in CVB3 (Verdaguer et al., 2003). VP2 This protein includes 263 residues, alternates with VP3 around the 3-fold axes of the virion and is relatively exposed at the capsid surface. VP2 encompasses a puff region composed of two sequential loops, one of them being more exposed on the surface of the particle (Fry et al., 2003). VP3 This protein contains 238 residues. Its N-terminal end is located close to the 5-fold axis of the virion being superimposed to the N-termini of VP1 and VP4. This structure together with those of the other 5-fold-related axes form a β-cylinder that is important for the stability of the pentamer. The C-terminal of VP3, on the other hand, is external and positioned adjacent to a major surface protrusion termed the ‘knob’ (Fry et al., 2003). VP4 This smaller protein, 69 residues in length, shows little secondary structure. It is the most conserved of the structural proteins and, as stated above, is located on the inner surface of the capsid. The glycine residue present at its N-terminus is covalently linked to a myristic acid group providing the capsid five symmetry-related myristoyl moieties around the inner surface of the capsid, a channel running from the inner to the outer surface at this point (Fry et al., 2003; Verdaguer et al., 2003). Rather than a continuous, circular canyon in SVDV there are five distinct depressions. The C-terminal of VP3, the first loop of the VP2 puff, and residues of the C-terminal VP1 loop form a ridge between these depressions (Fry et al., 2003). Non-structural proteins (NSPs) The P2 and P3 regions of the genome (Fig. 8.1) that encode the NSPs are highly conserved among enteroviruses, and the amino acid sequence of these proteins exhibits up to 90% homology. As mentioned above, SVDV encodes seven mature NSPs: 2Apro, 2B, 2C, 3A, 3B, 3Cpro and 3Dpol.

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2Apro This protease is responsible for polyprotein excision VP1/2A and the scission of the translation initiation factor eIF4GI within infected cells. eIF4GI cleavage contributes to the inhibition of the cap-dependent cellular protein synthesis, thus causing a shut off in host protein synthesis (Sakoda et al., 2001). Although most of the functions of the NSPs of SVDV have been inferred from those of the related enteroviruses, there is a number of studies pointing that 2Apro is important for the pathogenicity of this virus. In fact, the comparison between virulent and avirulent isolates suggested that key determinants of the pathogenicity in swine were located at the coding region for VP1/2A. Specifically, the presence of an arginine at the position 20 in the 2Apro is essential for induction of high viraemia and emergence of significant disease (Inoue et al., 2005, 2007). It has been proposed that, due to its ability to cleave eIF4GI, SDVD 2Apro regulates CAP-dependent protein synthesis but also can stimulate IRES driven viral protein synthesis (Sakoda et al., 2001). Accordingly, the modulation of these activities of 2Apro would be the responsible for the attenuation of SVDV. 2B, 2C and 3A These proteins accomplish functions not well characterized, but they may help viral replication by contributing to the proper replication complex organization, inhibition of cellular protein secretion, and virion assembly, as described for other picornaviruses (Greninger, 2015; Martín-Acebes et al., 2008; Vázquez-Calvo et al., 2016a). Supporting this view, these NSPs have been detected within viral replication complexes and colocalize with dsRNA intermediates that are markers for RNA replication (Martín-Acebes et al., 2008; VázquezCalvo et al., 2016a). Interestingly, enterovirus 2C exhibits a nucleotide helicase domain which could contribute to genome replication. 2C may also interact with host factors that regulate components of the cellular secretory pathway and control intracellular vesicle traffic, contributing to membrane rearrangements involved in replication complex assembly (Vázquez-Calvo et al., 2016a). Indeed, a single amino acid substitution in 2C protein (replacement of glutamine 65 for histidine) confers resistance to SVDV against treatment with Golgi disrupting agents (brefeldin A and golgicide) that impairs intracellular membrane traffic (Vázquez-Calvo et al., 2016a). 3B (VPg) This small protein is covalently linked to the 5′ end of the viral genome and is essential for virus replication being the uridylylated form the first step in this process (Schein et al., 2015; see also Chapter 3). 3Cpro This cysteine protease cleaves at certain Q/G sites in the viral polyprotein and, as described above, is responsible for most of the cleavages produced during polyprotein processing (Inoue et al., 1989). 3Dpol This protein is the viral RNA polymerase RNA dependent that is in charge of viral RNA replication and VPg urydylylation (Peersen, 2017; Sun et al., 2014). As commented in Chapter 3, the low copy fidelity inherent to this type of RNA polymerases as well as the vast

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size of virus populations give rise to population structures termed quasispecies comprising multitudes of related but non-identical individual genomes (mutant spectrum), subjected to continuous selection and sampling events (Domingo and Schuster, 2016). Antigenicity of the virion The antigenic structure of SVDV particles has been analysed using monoclonal antibody (MAb)-resistant (MAR) mutants (Kanno et al., 1995; Nijhar et al., 1999; Borrego et al., 2002a). These works defined seven neutralization sites highly dependent on conformation, involving amino acids not always contiguous in the protein sequence and even belonging to different proteins. When positioned on the corresponding capsid structure, these sites (named differently according to the different works) were found to be in four main areas, all of them well exposed on the surface of the capsid (Verdaguer et al., 2003) (Fig. 8.3). One of these areas is the BC loop of VP1, where amino acid residues VP1 83 and 84 were located. These residues contributed to define antigenic site 1, further dissected in sites Ia and Ib. Site Ia also involves residues VP1 95 and 98, in a location not so protruding (Borrego et al., 2002a). Another relevant area corresponds to the puff or EF loop in VP2, the largest loop of this protein. Most of the changes found in this protein map in this region, and two antigenic sites have been distinguished: site 2A involving positions VP2 160 and 163 within the second loop in the puff, and site 2B in the first loop (residues VP2 130 to 2154). The antigenic site 3 seems to be more complex in structure, involving residues mainly in VP3. Site 3A has been defined on a region including the VP3 ‘knob,’ which is – as mentioned above – the major surface protrusion of VP3, where residues 62 and 63 substituted in some

Figure 8.3  Antigenic sites involved in neutralization of SVDV. Outside view of a protomeric subunit of SVDV capsid highlighting the residues that have contributed to define each antigenic site (spheres). VP1 in blue, VP2 in green, VP3 in red, and VP4 in yellow. The positions of 5- and 3-fold symmetry axes are denoted by a pentagon or a triangle, respectively.

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mutants are located, together with the VP1 C-terminal residues 272 and 275. Site 3B consists of residues within the BC loop of VP3 and the C-terminal end of VP2 (residues 76 and 233). Site 3C is located at the carboxyl terminus of the protein (residues VP3 234 and 235). Finally, site IV, near the VP1 C-terminal end, is defined by amino acid 258. The MAb targeting this site is specific for isolates of the most recent antigenic group of SVDV that in addition can be associated with the ability of SVDV to bind HS (see next section). In addition to the characterization of MAR-mutants, other methods have allowed the identification of further antigenic regions in the SVDV capsid. The analysis by peptide scanning of a panel of sera from infected pigs led to the identification of novel antigenic regions recognized by SVDV-specific sera ( Jiménez-Clavero et al., 2000, 2001). One of these regions, the N-terminal end of the VP1 protein, was especially interesting. In contrast to the clear exposition on the virion surface of the sites identified by means of MAR-mutants, this site was only exposed and accessible to antibodies after the attachment of virus to cells, due to the conformational rearrangements occurred in the capsid upon binding to the cell receptor. A synthetic peptide spanning the 20 N-terminal residues of the VP1 capsid protein could efficiently mimic this linear site, since it was not only strongly recognized by antibodies from infected pigs, but was also able to induce antibodies capable to specifically block virus infectivity in vitro at some degree under certain conditions. Also by peptide scanning, but in this case analysing a panel of MAbs raised against the polyprotein P1, five main linear sites were identified (Borrego et al., 2002b). Although in some cases they overlapped in the areas already identified by MAR-mutants, none of the MAbs defining these sites were found to be able to neutralize viral infectivity, or to compete with the natural repertoire of antibodies induced by the infection, so their role in the antigenicity of the virus is still to be determined. Studies on the antigenic structure of SVDV have not only led to the identification and physical location of antigenic sites, but also revealed their level of conservation among SVD viruses and their variation in the field (Brocchi et al., 1997; Borrego et al., 2000; Bregoli et al., 2016). By analysis with a panel of MAbs representative of the different sites, the most stable site was proved to be site Ia, involving different amino acids in the region 83–98 of the VP1. No modification was detected in isolates of the four antigenic groups analysed, with the exception of minor changes in few strains from the last decade; furthermore, this site was also present in the human pathogen considered to be the ancestor of SVDV, coxsackievirus B5. Conversely, site Ib, defined by amino acids in positions 84 and 85 of VP1, in spite of its relationship with site Ia, was found to be more subject to variation. Monitoring of the antigenic profile of more than 200 Italian isolates collected over 25 years from 1992 showed that mutations in this site, initially sporadic, became fixed from 1997 for about ten years, whilst a probable retro-mutation, that caused reversion towards reactivity with the target MAbs, occurred again from 2008. In addition, site IV, stable until 2000, underwent changes evidenced by the reduced reactivity of the target MAb in more recent isolates (2006–2014), whilst antigenic modifications in other sites were sporadic and never became stabilized in the field (Fig. 8.3). These results are to be considered in order to design a proper diagnosis analysis based on MAbs directed to sites that may display different levels of conservation.

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Cellular biology of the virus Early infection steps: virus attachment, cell entry, and genome uncoating SVDV, like all other viruses belonging to the CVB5group, utilizes as cellular receptor a transmembrane glycoprotein of 46 kDa, known as CAR (coxsackie-adenovirus receptor) ( Jiménez-Clavero et al., 2005; Martino et al., 2000). The putative CAR binding site on the SVDV capsid has been predicted by overlaying the structure of CVB3–CAR with that of SVDV. This approach has revealed a moderate conservation of residues present at the CAR recognition site in both viruses as well as a similar distribution of hydrophobic and hydrophilic amino acids. The other surface molecule used as a co-receptor by some viruses of this group is DAF (decay accelerating factor, CD55), a 70 kDa protein also described to play an important role in the binding of SVDV (Martino et al., 2000). Interestingly, the use of DAF as co-receptor is only observed in early SVDV isolates ( Jiménez-Clavero et al., 2005), suggesting that as SVDV has been evolving in its new host, the pig, the structures of the capsid responsible for their binding to DAF have been lost. On the other hand, it has been reported the possibility that SVDV has acquired the use of another co-receptor, heparan sulfate (HS), since this glycosaminoglycan (GAG) has been shown to mediate the attachment of a more recent virus isolate, SPA’93, to host cells. Moreover, addition of soluble HS is able to completely inhibit infection so it cannot be ruled out the possibility that SVDV uses GAGs as an alternative receptor (Escribano-Romero et al., 2004). Whether HS replaces DAF as receptor in the pig, or participates in the cell binding as an alternative receptor or co-receptor, remains to be elucidated. The binding site of HS to SVDV has been identified through the analysis of heparin-resistant SVDV mutants (Escribano-Romero et al., 2004). Mutations in these viruses were located in the SVDV capsid structure revealing a HS binding site close to that identified in the FMDV–HS interaction (Verdaguer et al., 2003) and to the viral capsid region proposed by Fry et al. (2003) in the early isolates for their binding to DAF ( JiménezClavero et al., 2005). Thus it is possible that the alterations that the capsid has undergone in its evolution from CVB5 to SVDV are responsible not only for the loss of its binding to DAF but also for the increased binding to HS observed in recent isolates. The internalization of SVDV is dependent on clathrin-mediated endocytosis and plasma membrane cholesterol. Viral particles traffic through cellular endosomes and require specific cellular signalling and microtubule transport during these early infection steps (Martín-Acebes et al., 2009). Regarding the uncoating mechanism of the viral genome, it starts with virion binding to the cell surface. In fact, as commented previously, SVDV capsid undergoes a conformational change that allows the externalization of the N-terminus of VP1 ( Jiménez-Clavero et al., 2001). Thus, the current model hypothesizes that externalization of the N-terminus of VP1 is one of the first conformational rearrangements towards the production of the intracellular intermediates (also termed 35 S or A-particles) that precede viral uncoating (Bubeck et al., 2005; Butan et al., 2014). The blockage of this transition by binding of antibodies against this N-terminus of VP1 abolishes particle infectivity, as stated above ( Jiménez-Clavero et al., 2001). These intracellular uncoating intermediates should release VP4 and the viral genome to become empty particles. This process could take place within cellular endosomes (Martín-Acebes et al., 2009).

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Replication Once the viral RNA is released in the cytoplasm, it has to be translated to produce the structural and non-structural proteins that allow its replication and growth. Picornaviruses reorganize the cellular membranes of infected cells to generate specialized organelles for viral RNA replication. Translation and replication take place coupled to these membranous replication platforms via double-strand RNA (dsRNA) intermediates that are formed by the association of RNAs from positive and negative polarity. In enteroviruses, such as SVDV, RNA replication occurs at the surface of cytoplasmic vesicular structures derived from cellular membranes of the endoplasmic reticulum (ER) and the Golgi complex (MartínAcebes et al., 2008; Belov and Sztul, 2014; van der Linden et al., 2015). In the specific case of SVDV, infected cells undergo major ultrastructural rearrangements from 4 to 7 h post infection (Kubo et al., 1981). Confocal and transmission electron microscopy analyses have revealed that these rearrangements include disorganization of Golgi complex and ER that results in a proliferation of vesicular structures, including double membrane vesicles, within the cytoplasm of infected cells (Fig. 8.4). Such structures may provide the platforms for replication complex assembly (Martín-Acebes et al., 2008). As commented above, 2B, 2C, and 3A can induce these membrane rearrangements (Martín-Acebes et al., 2008; Greninger, 2015; Vázquez-Calvo et al., 2016a), albeit the mechanisms of membrane reorganization are not well understood yet. In some cases, the proposed mechanism of action of these NSPs involves their interaction with host cell proteins from the secretory pathway, which can vary among picornaviruses (Gazina et al., 2002; Martín-Acebes et al., 2008; van der Linden et al., 2010; Sasaki et al., 2012). In fact, the guanine nucleotide exchange factor (GEF) of the Arf family members of small GTPases involved in trafficking in the early secretory pathway GBF1 (Golgi-specific BFA resistance factor 1) could be localized in the viral replication factories of SVDV, as has been proposed for PV and coxsackievirus (Vázquez-Calvo et al., 2016a).

Figure 8.4  SVDV replication complex. (A) Confocal image of a IBRS-2 cell infected with SVDV. 2C protein (green) and dsRNA (red) were detected using adequate antibodies. The image was produced as described (Vazquez-Calvo et al., 2016a). Cell nucleus (blue) was stained using TO-PRO-3. (B) Transmission electron micrograph of an IBRS-2 cell infected with SVDV showing cytoplasmic accumulation of virus-induced vesicular structures. Cell were infected and processed for electron microscopy as described (Martín-Acebes et al., 2008).

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Virion assembly, maturation and egress Confocal analyses in infected cells have shown that both structural proteins and NSPs colocalize within SVDV replication complexes (labelled by 2C or dsRNA), suggesting that the assembly of capsid proteins is associated with the replication complex (Martín-Acebes et al., 2008). Cells infected with SVDV accumulate large amounts of virions within their cytoplasm that can even result in the formation of near crystalline arrays of viral particles (Kubo et al., 1980, 1981). After capsid assembly, VP0 is autocatalytically cleaved promoting virion maturation, and this cleavage is essential for virion infectivity (Rebel et al., 2003). Virions exit from the infected cell by cell lysis although in related enteroviruses release of viral particles can take place also via a non-lytic unconventional secretion mechanism related to the components of the cellular autophagic pathway (Bird et al., 2014). Viral pathogenesis SVD is a vesicular disease that can be subclinical or mild as well as severe (acute), but this latter form is usually associated with pigs housed on concrete flooring in humid conditions. The only known lesion directly attributable to the SVDV infection is vesicle formation on the coronary bands, on the heels of the feet and occasionally on the lips, tongue, snout and on the udder and teats in lactating sows. The severity of the clinical signs is dependent on flooring type – severe on concrete, mild on straw. In severe cases, the horn of the hoof sloughs off. Fever is usually absent and mortality is rare (Lahellec and Gourreau, 1975). Despite its low severity of infection, the lesions that SVDV produces are indistinguishable from those of FMD and other vesicular diseases in pigs. Indeed, as occurred in Taiwan in 1997, SVD can mask FMD outbreaks. Thus, the epidemiological importance of SVD is that the clinical signs of this economically unimportant disease are indistinguishable from those of FMD, which is an economic disaster if it occurs in a given area or country. Therefore, laboratory analyses play a crucial role in order to discriminate between the different viruses producing identical pathology (see Diagnostic procedures). Diagnostic procedures The World Organization for Animal Health (OIE) and the European Commission Decision 2000/428/EC regulate the requirements for SVDV diagnosis, concerning both the nature of samples to be analysed as well as the kind of tests to be performed. The selection criteria have to consider the timing of infection, the clinical or subclinical occurrence and the tests’ sensitivity. Infected animals go for a period of 2–7 days of incubation before the appearing of lesions. However, up to 48 hours before the onset of clinical signs virus may be excreted from nose and mouth of the affected pigs and also in the faeces. This viral production continues for the first 7 days after infection, and usually ends within 2 weeks. Virus excretion in the faeces may continue up to 3 months. Thus, epithelium and oral fluids when the disease is clinically evident, or faecal samples to investigate both clinical and subclinical suspects can be used for diagnoses. Classical techniques focused on the identification of the agent, either viral particles by ELISA or viral genome by RT-PCR, are suitable screening tests, rapid and easy to perform. However, ELISA suffers for lower sensitivity and can therefore be applied only with virus rich samples, like vesicular epithelium. Viral isolation on cell culture is the

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reference method, however it requires expensive and specialized equipment and at least 6 days before results are available for negative samples. Only samples collected early after infection will allow the detection the viral agent whereas serological tests performed with samples collected later will be able to confirm that infection, including subclinical infection, occurred. The identification of the isotype of antibody in positive samples will also allow discriminating between recent infections indicative of virus shedding (IgMs alone or together with IgGs) and older exposure to infections (only IgGs) (World Organisation for Animal Health, 2018). Diagnoses of SVDV is sometimes problematic by the phenomenon called ‘singleton reactors’: single animals within a cohort that even although never being exposed to the virus, may react positively in serological tests for antibody to SVD virus, both ELISA and even by the VN test, while the other animals remain negative. These ‘singleton reactors’ constitute a small proportion, less than 1%. Infection by other viruses or other non-specific serum factors may be responsible of the serological cross-reactivity, but the factors remain unknown. A variety of RT-PCR methods have been developed for the detection of SVDV both conventional (Vangrysperre and De Clercq, 1996; Lin et al., 1997; Núñez et al., 1998; Callens and De Clercq, 1999) and in real-time format (Reid et al., 2004; McMenamy et al., 2011), employing different techniques for RNA extraction and targeting different parts of the SVD virus genome, some of them also aimed to the differentiation of SVDV from FMDV and VSV (Hakhverdyan et al., 2006). A one-step reverse transcriptase loop-mediated isothermal amplification (RT-LAMP) assay has also been developed (Blomstrom et al., 2008). However, only a few of these methods have been fully validated. The conventional RT-PCR (Núñez et al., 1998) combined with an RNA extraction method based on immuno-purification of the virus (Fallacara et al., 2000; World Organisation for Animal Health, 2018) succeeded in SVDV control and eradication in Italy, where the national surveillance plan consisted in testing faecal samples for the presence of SVDV. In a comparative study conducted by Benedetti et al. 2010 on positive faecal samples from many different SVD outbreaks occurred in Italy, the conventional one step RT-PCR, targeted on the 3D gene, showed the best diagnostic performance, with the capability to reveal all the circulating genomic sublineages, with respect to two real-time RT-PCR assays targeting the 5′-untranslated region (Reid et al., 2004), and an RT-LAMP (Blomstrom et al., 2008). SVD virus is usually present in low concentration in faeces, thus in order to succeed in virus detection, a high sensitive method, less affected by mutations present in the primer/ probe target sequence, is needed. Among real-time RT-PCR assays the one targeting the 5′-untranslated region named 2B-IR (Reid et al., 2004) showed less diagnostic sensitivity on faecal samples in respect to the conventional assay (Benedetti et al., 2010), due to primers mismatching and the low amount of virus present, however it was able to detect the same virus after isolation on cell culture. On the contrary, Real-Time RT-PCR based on 3-IR primers/probe set (Reid et al., 2004) and RT-LAMP were unable to amplify genomes of one sublineage circulating even after cell culture isolation, for the presence of mutation within the probe target sequence for 3-IR and within primers target sequence for LAMP. New molecular methods are being developed to improve the detection capability of those already available. Features searched for these novel assays are the speed for detection, the ability to simultaneously detect and identify different swine viruses and the portability of the devices.

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Epidemiology Transmission Swine is the sole natural host known for SVDV. The virus does not cause death and morbidity rate in herds may be low but high in groups of pigs (in pens). The virus infects swine via lesions in skin and mucosa, ingestion and inhalation, and low amounts of virus are required to infect animals across broken skin. Affected pigs may excrete virus from the nose and mouth and in the faeces up to 48 hours before the onset of clinical signs and all tissues contain virus during the viraemic period. SVDV is stable in a wide pH range and therefore it is not inactivated by normal pH change associated with rigor mortis. Most virus is produced in the first seven days after infection, although virus excretion may be observed up to three months in the faeces. Ruptured vesicles (epithelium and fluid) are a high-titre source of virus, while faeces are a lower-titre source of virus. Direct contact among infected swine or with their excretions are major sources of virus spread, often within contaminated vehicles or premises. As detailed in the OIE Terrestrial Animal Health Code, movement of subclinically infected animals is considered the most common means of spreading SVDV. Transport of large numbers of swine often results in small lesions that provide a portal of entry for the virus. Non-heat-treated garbage fed to swine provides another means for infected meat to cause disease. Occurrence and history In Europe SVD is reported only occasionally, except in Italy, where the virus has been detected until 2015. In this country continuous surveillance has been in place since 1995, aimed at monitoring subclinical virus circulation, through virus detection in faeces and antibody detection in sera, according sampling schedules laid down in a national surveillance and eradication plan. Italy experienced SVD for the first time in 1966, and during almost 50 years of permanence four main viral antigenic variants have been described (Brocchi et al., 1997): the first variant corresponds to the first virus isolated in 1966 in Italy, the second includes viruses circulated in Europe and Far East during the 1970s and 1980s, the third includes viruses circulating from 1988 to 1992 only in Italy and the fourth variant persisted from 1992 to 2015. Interestingly, outbreaks caused by this last variant were first reported in The Netherlands (1992) and then sporadically also in Spain and Portugal. Phylogenetically, the four antigenic variants are distributed in four distinct genetic clusters. The availability of all isolates collected during 25 years of circulation in Italy and analyses based on full genome sequences enabled the study of the molecular evolution of viruses of the most recent variant, within which two main sublineages can be distinguished, both derived from a unique common ancestor dated back to 1990–91. Viruses of the sublineage named A are organized in two distinct groups: one is composed of viruses circulated in Italy from 1992 to 1995 and closely linked to isolates detected in The Netherlands (1992 and 1994) and Spain (1993); a second group, that includes Italian and Portuguese isolates, is on its turn divided in two distinguishable clades, temporarily separated but with a common origin. The long temporal distance and the low genetic divergence between these two clades suggest that the virus of the older clade (1995–1999) has been

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circulating undetected and resurged after many years either in Portugal (in 2003) or in Italy (in 2004) (Knowles et al., 2007), where it spread in central-southern regions up to 2010. The sublineage named B evolved only in Italy from 1993 to 2010. After a period of co-circulation in northern and southern Italy, from 2000 this sublineage was maintained in southern Italy causing almost exclusively subclinical forms, with sporadic incursion in central-northern regions, which were promptly eradicated. The last incursion in North Italy occurred in 2006–07 with two distinct epidemic waves (Bellini et al., 2010). From 2007 to 2010 strains originated from recombination between lineages A and B co-circulated with parental strains, later on the recombinants remained the unique variant still detected (manuscript under preparation). Prevention and control of the disease: vaccination Control measures set out by the international legislation (OIE) against SVD have been always restricted to sanitary prophylaxis. These measures, which are based on surveillance programmes, are elimination of infected and contact pigs (stamping out), movement controls and quarantines for animals and animal products, strict import requirements and disinfection of premises, transport vehicles and equipment. Disinfection is extremely important due to the physical properties of the virion: very stable in the environment and resistant to pH 2–12. A good example is provided by the measures taken in an outbreak of the disease which occurred in Italy in 2007 (Bellini et al., 2010; Nassuato et al., 2013). Prophylactic (vaccines) or therapeutic (antivirals) interventions have never been considered as actions to control SVD. Indeed, to our knowledge, no testing of compounds as therapeutic antiviral candidates against SVDV have been reported in the literature, and actually there are no treatments or commercial vaccines currently available against the disease, even although some experiments have analysed the ability of different formulations to induce protective immunity. In the 1970s, when the disease was dramatically affecting Europe, a few works analysed classical formulations based on chemically inactivated virus. Although the studies were not exhaustive, results revealed that it was possible to induce protection against viral infection in the vaccinated animals (Lahellec and Gourreau, 1975; Mitev et al., 1978). Further studies following important outbreaks in Europe and Asia in the 1990s analysed the immune responses induced by subunit vaccines produced by recombinant technology. As already done for other picornaviruses such as FMDV, the antigen selected as immunogen was mainly the SVDV capsid precursor polypeptide (P1), presented in different vaccine formulations (see Chapter 3). Recombinant P1 produced in bacteria did not induce a very efficient immunization after a single injection in swine, the natural host of the virus. Even although animals developed both humoral and cellular responses, no neutralizing antibodies were detected ( Jiménez-Clavero et al., 1998), probably because conformational epitopes critical for the recognition of neutralizing antibodies are not well mimicked in these constructs. More recently, a suicidal DNA vaccine based on the alphavirus replicon, supposed to present the P1 polypeptide in a manner more similar to that of a natural infection, was assayed in guinea pigs and swine (Sun et al., 2007). Even although four out of six pigs developed neutralizing antibodies after receiving three intramuscular injections of the DNA vaccine, when challenged with live SVDV only 50% (3/6) of the animals were protected. Thus, a lot of work remains to be done in order to develop an efficient vaccine able to protect against SVDV infection, including also immunological studies on the immune correlates

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that allow the identification of those parameters of the immune response that are important for protection against viral replication and disease. Such findings could maybe lead to some change in the control measures established by international legislation. References

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Index

A

D

Ad5 65 Adaptation 22, 23, 49, 62, 83, 109, 124, 138, 181–184 Adjuvants 11, 62, 142, 166 Antibody 4, 9, 24, 25, 29, 31, 51, 55, 56, 93, 96, 98, 118, 119, 125, 142, 157, 162, 164, 166–168, 187, 192, 193 Antigenic variants 59, 62, 193 Antigen presentation 8, 55 Antigen-presenting cells 167 Antiviral agent 12, 13 Antiviral drug 2, 12, 13, 126 Apigenin 13 Assembly 5, 6, 23, 46, 47, 50, 112, 151, 154, 158, 183, 186, 190, 191 Attenuated 10, 11, 21, 32, 33, 47, 62, 63, 112, 114, 124, 164 Autophagy 6, 7, 23, 157, 160

Dendrimer peptide 65 Dendritic cell 24, 55, 91, 94, 156, 161 Diagnosis 1, 9, 30–32, 43, 59, 60, 81, 86, 87, 90, 95–97, 108, 118, 123, 135, 140, 141, 162, 181, 182, 188, 191 Diagnostic test 10, 22, 30, 31 DIVA (differentiating infected from vaccinated animals) 31–33, 62–64, 96, 98, 165, 166 DNA replication 5, 6, 13, 136,

B B-cell 64, 65, 92, 168 Biosecurity 81, 98, 115, 120, 122, 123, 125 B-lymphocyte 8, 28, 29

C Capsid 3, 43, 44, 46, 47, 50–52, 55, 56, 60, 63, 64, 81–84, 93, 97, 98, 109, 136, 137, 142, 150, 165, 182–185, 187–189, 191, 194 Capsomer 3 Cell culture adaptation 22, 124 Clinical sign 7, 9, 11, 24–26, 28, 43, 58, 59, 81, 83, 86–91, 96, 98, 116, 123, 135, 138, 140, 144, 160–162, 166, 181, 182, 191, 193 Compartment 55, 113 Conventional vaccine 62, 65 Cytokine 9, 21, 28, 29, 52, 65, 85, 92, 94, 114, 160, 161, 167 Cytotoxic 63, 161 Cytotoxic T-cell 29, 55, 92, 162

E ELISA (enzyme-linked immunosorbent assay) 9, 10, 13, 31, 60, 96, 118, 119, 141, 162, 191, 192 Empty capsid 47, 63, 183 Encapsidation 47, 156, 183 Endemic 2, 21, 32, 61, 62, 115, 118, 121, 122, 125, 135, 141, 162, 163 Endocytosis 5, 22, 23, 56, 137, 154, 156, 189 Epidemics 107, 111, 115, 120–125 Epidemiology 1, 23, 24, 31, 43, 61, 62, 82, 95, 108, 111, 119–121, 141, 149, 163, 169, 182, 193 Epitope 51, 55, 64, 65, 93, 119, 164, 165, 167, 168, 194 Eradication 32, 61, 192, 193 Evolution 33, 44, 53, 82, 83, 98, 113, 121, 122, 138, 143, 155, 189, 193

F FAO (Food and Agriculture Organization of the United Nations) 60 Flavonoid 13

G Genistein 13 Genome organization 43, 84, 85, 108, 110, 151, 181, 182 Guinea pig 32, 48, 62, 64, 140, 194

200  | Index

H Helicase 23, 48, 53, 57, 58, 85, 110, 152, 153, 155, 186 Heparan sulfate 56, 84, 112, 189 Host cell 4, 5, 13, 22, 46, 47, 52, 53, 57, 149, 151, 154, 157–159, 164, 169, 189, 190 Host range 48, 58, 112 Humoral 28, 29, 52, 55, 59, 92, 93, 161, 194

I Immune response 8, 11, 12, 23, 24, 28, 29, 52–55, 59, 63, 65, 86, 91–93, 125, 139, 142, 155, 157, 158, 161, 164, 166, 167, 194, 195 Immunomodulation 95 Innate immunity 53, 114 Integrin 56 Interferon 23, 28, 47, 54, 58, 65, 85, 94, 112, 114, 157–159, 161, 162 Internal ribosome entry site (IRES) 44, 45, 182 Isotype 192

Pathogenesis 7, 21, 24, 28, 43, 56, 58, 88, 89, 92, 93, 108, 112–115, 117, 135, 149, 158, 160, 181, 191 Pentamer 50, 51, 57, 62, 63, 183–185 Persistence 24, 25, 30, 59, 61 Phylogenetic 3, 23, 39, 61, 83, 105, 110, 111, 120–122, 124, 127, 138, 143, 147, 182, 193 Polymorphism 95, 161 Polyprotein 6, 22, 44, 46, 47, 49, 53, 63, 109, 110, 113, 151, 152, 182, 183, 186, 188 Protection 10, 11, 29, 33, 52, 55, 60, 63–65, 93, 97, 98, 119, 124–126, 142, 164–169, 194, 195

Q Quasispecies 49, 187

R

Lethal dose 11 Lethal mutagenesis 65 Livestock 43, 61, 64, 65

Real-time PCR 9, 10, 30, 118, 192 Receptor binding 23, 63, 119, 137 Recombination 81–83, 98, 110, 121, 122, 165, 194 Replication complex 47, 48, 58, 152, 186, 190, 191 Resistance 1, 10, 47, 64, 98, 117, 119, 126, 141, 186 Ribavirin 65, 126 RNA genome 22, 110, 151, 152, 156 RNA replication 47–49, 186, 190

M

S

K Knockout 4, 156

L

Macrophage 4–9, 11–13, 24, 28, 29, 58, 89–92, 94, 112, 137, 156, 160–162, 166 Monoclonal antibody 51, 118, 187 Mutant 7, 11, 48, 51, 52, 64, 138, 141, 166, 187–189 Mutation 11, 33, 44, 48 49, 56, 63, 81, 83, 98, 109, 122, 142, 155, 164, 188, 189, 192

N Negative-strand RNA 109, 152, 154 Neutralizing antibodies 4, 29, 55, 63, 64, 109, 119, 125, 162, 164, 194 Next-generation sequencing 85, 86, 95 Nucleoside analogue 12, 65

O OIE (World Organization for Animal Health) 21, 23, 30, 60, 61, 191, 193, 194 Opsonization 55 Outbreak 2, 10, 24, 31, 59–62, 65, 82, 107, 108, 111, 115, 118–122, 124–126, 162, 191–194

P Passive immunity 162 Pathogen-associated molecular patterns (PAMPs) 52

Sequencing 2, 23, 60–62, 85, 86, 95, 111, 118, 182 Serotype 23, 44, 51, 52, 60–62, 64, 110 siRNA 13 Strain 1, 2, 4, 7–11, 23–25, 27, 28, 31–33, 81–83, 91, 93, 98, 108–112, 114, 120–122, 124, 125, 135, 138, 142, 150, 160–167, 169, 182, 188, 194 Surveillance 32, 120, 192–194 Synthetic peptide 64, 96, 167, 188

T T-cell 28, 29, 55, 64, 65, 92, 93 T-helper 8, 29 T-lymphocyte 8, 28, 29, 55, 92 Toll-like receptor (TLR) 52, 158 Transcription factor 6, 53, 54, 158 Transmission 2, 31–33, 60, 61, 81, 82, 91, 98, 115, 123, 126, 142, 162–164, 190, 193

U Uncoating 56, 57, 84, 112, 113, 154, 185, 189

V Vaccination 11, 21, 31–33, 43, 55, 61, 62, 64, 65, 81, 93, 97, 98, 115, 124, 125, 135, 141, 142, 162, 164, 166, 181, 194 Vaccine strain 112, 121, 125, 142, 164, 165 Variability 4, 23, 24, 62, 161, 182

Viraemia 7, 32, 58, 89, 92, 137, 142, 160, 164, 166, 186 Virulence 1, 4, 7, 11, 22–25, 28, 46, 49, 56, 62–64, 82, 83, 86, 112, 137, 149, 150, 163–165

W Wild boar 1, 21, 26, 28, 31–33

Porcine Viruses From Pathogenesis to Strategies for Control The global population has quadrupled over the last century leading to an increased demand for affordable safe food. Satisfying this demand will not be easy and will require even more widespread use of intensive farming practices. However, intensive farming practices can lead to higher probabilities of outbreaks of a variety of viral diseases, a critical concern in terms of food protection and food security. In the case of the pig industry there are several important viruses. One example is African swine fever virus, which causes a devastating disease with enormous socio-economic consequences in affected countries, mostly in Africa. No African swine fever virus vaccine currently exists. An understanding of the molecular biology, pathogenesis, host–virus interactions and epidemiology of these viruses is critical for their prevention and control. This book provides a comprehensive review of the current knowledge of the most important porcine viruses, written by prominent scientists who have made great contributions in their respective fields of expertise. Topics include African swine fever virus, classical swine fever virus, foot-and-mouth disease virus, porcine circoviruses, porcine epidemic diarrhoea virus, porcine parvovirus, porcine reproductive and respiratory syndrome virus and swine vesicular disease virus. Each chapter covers the current knowledge on epidemiology, pathogenesis, virus biology, diagnosis, and prevention and control strategies. This book is essential reading for everyone working with porcine viruses, from PhD students to experienced scientists, in academia, the pharmaceutical or biotechnology industries and clinical environments.

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