Development of an Improved Live Attenuated Antigenic Marker CSF

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9-9-2016 Development of an Improved Live Attenuated Antigenic Marker CSF Vaccine Strain Candidate with an Increased Genetic Stability lauren Gayle Holinka-Patterson [email protected]

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Development of an Improved Live Attenuated Antigenic Marker CSF Vaccine

Strain Candidate with an Increased Genetic Stability

Lauren Holinka-Patterson, PhD

University of Connecticut, 2016

Abstract

Classical Swine Fever Virus (CSFV) is an extremely contagious, hemorrhagic and often fatal disease of pigs that causes serious socioeconomic impact on countries which experience outbreaks or are endemically infected. Current control measures utilized for CSFV are contingent upon the epidemiological status of the inflicted area and entail either prophylactic vaccination or non-vaccination, “stamping out”, protocol with the elimination of infected herds and culling of animals on neighboring farms. This latter practice results in less than satisfactory consequences, although once thought to be the answer to eradication of CSFV from a region,

“stamping out” results in tremendous economic losses as well as the ethical objection to the potential killing of millions of healthy pigs. Marker vaccines also referred to as DIVA vaccines because they allow for differentiation between naturally infected and vaccinated animals

(DIVA), could complement or replace the “stamping out” strategy. The use of a DIVA vaccine with an accompanying diagnostic test would make mass vaccination of susceptible pigs during and outbreak possible thereby preventing the rapid spread of the virus while allowing for identification of CSFV infected animal through serological surveillance. With the advent of reverse genetic technology, it is now possible to rationally design a CSFV vaccine that contains such markers. Recently, we reported the development of a live attenuated CSFV strain with two antigenic markers, named Flag T4v. During the vaccine assessment process, Flag T4v showed

evidence of reverting back to the virulent phenotype of wild type CSFV. Analysis of the genome sequence from the recovered revertant virus revealed the presence of four non synonymous substitutions and a deletion of one of the antigenic epitopes compared to the parental Flag T4v genome. In order to improve the genetic stability of Flag T4v, the nucleotide codon sequence in these regions was modified as much as possible from its original sequence without compromising the wild type amino acid sequence by taking advantage of codon redundancy in an attempt to make viral reversion more difficult while maintaining both viral attenuation and reactivity of the antigenic markers. The newly developed virus, Flag T4Gv, was shown to be stably attenuated when assessed in a reversion to virulence test. In addition, Flag T4Gv was also shown to possess similar efficacy of Flag T4v in terms of protecting swine at early and late times post vaccination.

Development of an Improved Live Attenuated Antigenic Marker CSF Vaccine

Strain Candidate with an Increased Genetic Stability

Lauren Holinka-Patterson

B.A., University of Connecticut, 2001

M.A., University of Quinnipiac, 2007

A Dissertation

Submitted in Partial Fulfillment of the

Requirements for the Degree of Doctor of Philosophy

at the

University of Connecticut

2016

Lauren Holinka-Patterson

2016

APPROVAL PAGE

Doctor of Philosophy Dissertation

Development of an Improved Live Attenuated Antigenic Marker CSF Vaccine Strain Candidate

with an Increased Genetic Stability

Presented by

Lauren Holinka-Patterson, B.A., M.A.

Major Advisor______Guillermo Risatti

Associate Advisor______Manuel Borca

Associate Advisor______Paulo Verardi

University of Connecticut 2016

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ACKNOWLEDGEMENTS

Over the past fourteen years it has been my great pleasure to be a member of the Foreign

Animal Disease Research Unit (FADRU) at Plum Island Animal Disease Center (PIADC). My supervisor and committee member, Dr. Manuel Borca, has been a great inspiration and exceptional mentor to me. His innovative thinking has made him a leader in the study of CSFV,

ASFV and FMDV worldwide. I am grateful for the guidance, support and encouragement he has provided me over the course of this work and throughout my career.

I would also like to thank my major advisor, Dr. Guillermo Risatti for sharing his tremendous wealth of knowledge in the areas of microbiology, virology, and pathobiology. I especially would like to thank Dr. Risatti for his ability to explain things clearly and simply when I needed it most. I am also grateful to committee member, Dr. Paulo Verardi for his support and advice along the way.

I am also very appreciative for all the assistance that I received from Dr. Zhiqiang Lu with the sequencing of the mutant DNA plasmids and rescued viruses. Lastly, I would like to acknowledge and extend my gratitude to the animal handlers at PIADC for the outstanding service that they provide our scientific staff on a daily basis.

Finally, I would like to express my gratitude to my family and friends for their unwavering love and support throughout this work.

TABLE OF CONTENTS

Page

INTRODUCTION ...... 1

Literature Review ...... 4

Genome ...... 4

5’UTR ...... 5

3’UTR ...... 5

Npro ...... 6

Core ...... 8

Erns ...... 13

E1 ...... 17

E2 ...... 20

p7 ...... 27

NS2 ...... 29

NS2-3 ...... 30

NS3 ...... 31

NS4A ...... 33

NS4B ...... 33

NS5A ...... 36

NS5B ...... 37

Viral Life Cycle ...... 38

Translation and Protein Processing ...... 39

RNA Replication...... 41

Clinical Signs...... 42

Pathogenesis...... 45

Viral Transmission...... 47

Epidemiology...... 47

CSFV Reintroduction Pathways into the US ...... 49

Intentional Release ...... 49

Unintentional Release ...... 50

CSF Control Strategies...... 52

Live Viral Vector Vaccine Candidates...... 55

Non-replicating Subunit Vaccine Candidate ...... 57

Recombinant Pestivirus Vaccine Candidates ...... 59

CP7_E2alf Marker Vaccine Candidate ...... 71

Flag T4v Marker Vaccine Candidate ...... 76

Thesis Research ...... 78

MATERIALS AND METHODS ...... 81

RESULTS...... 97

DISCUSSION...... 110

REFERENCES...... 118

LIST OF FIGURES

Page

Figure 1. Schematic representation of CSFV polyprotein ...... 138

Figure 2. Schematic representation CSFV Virion Structure...... 139

Figure 3. Global distribution of CSFV in 2014 ...... 140

Figure 4. Schematic representation CSFV RB-22v and C22-Flagv ...... 141

Figure 5. Schematic representation of CSFV T4v polyprotein ...... 142

Figure 6. Schematic representation of CSFV Flag T4v polyprotein ...... 143

Figure 7. Daily temperature of animal groups in minimum protective dose study ...... 144

Figure 8. Depiction illustrating altered codon usage for Flag T4G ...... 145

Figure 9. Growth Curve of Flag T4Gv compared with Flag T4v and BICv ...... 146

Figure 10. Antigenic profile of Flag T4Gv ...... 147

Figure 11. Viremia Flag T4Gv protection assessment experiment ...... 148

Figure 12. Viremia of Flag T4GV Early Protection Study ...... 149

Figure 13. Cytokine Quantification in Serum...... 150

Figure 14. qRT-PCR Cytokine Expression ...... 151

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LIST OF TABLES

Page

Table 1. Amino acid and nucleotide changes observed in revertant Flag T4SPv...... 152

Table 2. Nucleotide sequence of primers used for sequencing cDNA IC...... 153

Table 3. Nucleotide sequence of primers used for evaluation of Flag T4SPv ...... 156

Table 4. Nucleotide sequence of primers for Flag T4SPv PCR amplification ...... 157

Table 5. Determination of minimum protective dose experiment ...... 158

Table 6. Survival and fever response in pigs infected with CSFV chimeric viruses representing mutations in Flag T4SPv ...... 159

Table 7. Nucleotide sequence of primers used in production of Flag T4Gv ...... 160

Table 8. Swine survival and fever response in Flag T4Gv infected animals after challenge with parental virulent BICv ...... 161

Table 9. Swine survival and fever response in Flag T4Gv infected animals challenged with BICv at different time points post infection ...... 162

Table 10. Primer Sets for qRT-PCR Cytokine Gene Expression Assay...... 163

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INTRODUCTION

Classical swine fever (CSF) is an extremely infectious and generally lethal hemorrhagic disease of domestic and wild pigs. The causative agent, classical swine fever virus (CSFV), is a small enveloped virus with a single stranded RNA genome with positive polarity. It belongs to the genus pestivirus within the family Flaviviridae (Francki et al., 1991). The 12.5 Kb CSFV genome contains a single open reading frame that encodes a 3898-amino-acid polyprotein which is processed co- and post translationally into 11 to 12 proteins (NH2-Npro-C-Erns-E1-E2-p7-

NS2-NS3-NS4ANS4B-NS5A-NS5B-COOH) (Rice, 1996). Structural components of the CSFV virion include the capsid (C) protein and glycoproteins Erns, E1, and E2. E2 is the most immunogenic of the CSFV glycoproteins (Weiland et al., 1990), inducing neutralizing antibodies, which provide protection against lethal CSFV challenge.

CSFV is endemic in many parts of the world, including Asia, Africa, South and Central

America. Outbreaks have also been reported in the Caribbean nations of Cuba, Haiti and the

Dominican Republic. In Europe disease outbreaks occur intermittently due to the endemic presence of the CSFV in the wild boar population. Regions currently believed to be free of CSF include Australia, New Zealand, Scandinavia, Iceland, Ireland, Canada and United States. The last reported case of CSFV in the United States was in 1976 with official CSFV free status recognized in 1978 after a 16-year effort by the farm industry and State and Federal governments.

Due to the inherent ability of CSFV outbreaks to cause extreme detrimental socioeconomic consequences the Office International des Epizooties (OIE) has classified CSFV as a notifiable list A pathogen. The OIE is ultimately responsible for setting outlines for internationally agreed upon standards relating to animal health and conditions under which a country can be considered free of CSF which consequently affects the ability of a country to engage in international trade of pigs and pig products. Due to the highly infectious nature of

CSF, disease free countries have imposed trade barriers that prohibit importation of animals or animal products from countries that do not possess CSF free status with the ban continuing until the OIE recertifies that country as CSF free.

Although the United States has been free of classical swine fever (CSF) since 1978, today’s ease of global mobility has increased the possibility of livestock diseases to travel the world which has raised concerns about how an outbreak of CSFV would affect the United States.

Agriculture is the largest employer and industry in the United States, generating more than $1 trillion annually (DHS, 2008). A major component of this is livestock, which contributes over

$100 billion annually to the gross domestic product. The introduction of CSFV at the farm level would cause severe economic disruption given that agriculture accounts for 13 percent of the

U.S. gross domestic products and 18 percent of domestic employment (DHS, 2008). This economic disruption could potentially be exacerbated by long-lasting trade embargoes imposed on U.S. pork and pork products.

Currently, there are two main strategies utilized in the prevention and control of CSFV, prophylactic vaccination or non-vaccination “stamping out” policy with control strategy

selection dependent upon the epidemiological status of the country. Prophylactic vaccination with traditional live attenuated vaccines (LAVs) is commonly employed in regions where CSF is endemic, such as, Asia, Africa, South and Central America. However, the humoral immune response induced by these vaccines does not differ from that elicited by infections caused by the wild type viruses as such vaccinated animals cannot be serologically differentiated from infected animals. As a result, once emergency vaccination is implemented in a region, that region is thereafter prohibited from international trade for a period of at least 2 years from the last reported case of CSFV (van Oirschot et al., 2003). The resulting ban on pork and pork products result in major economic losses for the affected region. The second strategy for CSF control is a non- immunization “stamping out” policy which has a more favorable time frame in the return to CSF free status. Due to the economic benefits of rapid return to disease-free status, the European

Union (EU) and some parts of Eastern Europe have opted for this latter approach. However, the large scale culling of animals resulting from this practice is very problematic in that many healthy animals eventually have to be destroyed for welfare reasons. Due to the huge economical losses caused by the “stamping out” policy as well as ethical objections to the mass extermination of millions of healthy animals, alternatives to the current policies are needed.

One important option being considered which suitably satisfies this problem is the development of safe LAV “marker” or differentiating infected from vaccinated animal (DIVA) vaccines, which are intended to be used with a companion diagnostic test which makes differentiation of infected animals from vaccinated animals possible (van Oirschot et al., 2003).

The use of this type of vaccine would limit loss of livestock impacting the overall economic ramifications but could also be potentially used for CSF eradication. Development of such a

vaccine is of the utmost importance to the US and nations of the European Union which do not currently have a CSFV control plan that includes emergency vaccination. As such, the purpose of this study is to develop a safe and effective marker or DIVA vaccine to fulfill this need in the event that CSFV was reintroduced either intentionally or unintentionally into the US. The solution presented here is the development of a rationally designed safe and effective LAV with

DIVA capabilities using reverse genetic technology.

Literature Review

Genome

Pestiviruses are small spherical-enveloped viruses approximately 40-60 nm in diameter

(Fig. 1). The envelope surrounds an electron-dense hexagonally-shaped nucleocapsid that is about 30nm in diameter (Wengler et al., 1995). The nucleocapsid contains a single-stranded

RNA genome of positive polarity with an approximate length of 12.5kb. The viral genomic

RNA is neither capped nor polyadenylated and contains one large open reading frame (ORF) coding for a polyprotein of approximately 4000 amino acids flanked by a 5’ and 3’ untranslated region (UTR). The polyprotein is processed co- and post translationally by viral and host cell proteases producing either 11 or 12 mature proteins. The order of the cleaved products is as follows: NH2-Npro-C-E0-E1-E2-p7-NS2-3(NS2, NS3)-NS4A-NS4B-NS5A-NS5B-COOH (Fig.

2).

5’UTR

The 5’ UTR is comprised of cis-acting elements that are believed to be involved in both viral replication and protein expression. The first of these elements is a stem loop called the Ia hairpin which is composed of approximately the first 30 nucleotides of the viral sequence.

Experiments with a replicon of the closely related pestivirus, BVDV, determined that the Ia hairpin is involved in both translation initiation and replication of pestivirus RNA with the first four residues of the genome necessary for replication (Grassmann et al., 2005; Yu et al., 2000).

Pestivirus RNAs lack 5’cap structure which normally promotes translation initiation (Brock et al.

1992). Instead, pestiviral RNAs contain an internal ribosomal entry site (IRES) that is able to directly attract the small ribosomal subunit and position it at the translational start site (Poole et al., 1995). The CSFV IRES belongs to the hepatitis C virus/pestivirus (HP) group of IRES which are characterized by two complex stem-loop structures denoted as domains II and III. The initiation site is located downstream of domain III with a pseudoknot located between them. This pseudoknot is essential for IRES function as it is crucial for orienting the AUG start codon in the

P-site of the ribosome (Moes et al., 2007).

3’ UTR

Similar to the 5’UTR, the 3’UTR of CSFV is also believed to contain important cis- acting elements. The 3’UTR is able to fold into three stem loops designated SLI, SLII, and SLIII.

SLI is located closest to the 3’ end of the viral sequence, while SLII is positioned between SLI and SLIII, with SLIII directly downstream of the stop codon. The SLI loop has been shown to be

indispensible for pestiviral replication by functional analysis by both changing the codon usage of SLI which maintained the conserved secondary structure and by changes to the SLI nucleotides sequence which disrupted the naïve stem loop (Yu et al., 1999). Viable mutants were only obtained where the SLI loop remained unaltered. Surprisingly, pestiviral RNA do not contain a Poly(A) tail common in most eukaryotic mRNAs which is known to influence RNA stability as well as function to support translation initiation (Hinnebusch et al., 2012).

Npro

The first protein encoded for by the CSFV genome is a nonstructural protein, termed

Npro, which possesses autoprotease activity that enables the protein to cleave itself at its C- terminus from the downstream nucleocapsid protein (Tratschin et al., 1998). Interestingly, while the other two members in the Flaviviridae family (flaviviruses and hepatitis C virus) have similar overall genome organization as the pestiviruses, neither contains an analogous counterpart to

Npro.

Protein crystallization experiments by two separate groups revealed that the Npro structure forms a novel fold characterized as a “clamshell” fold. (Gottipati et al., 2013; Zogg et al., 2013). Npro is composed of mostly β-sheets and can be divided into two distinct domains, the catalytic protease domain and the zinc-binding domain. The protease domain contains coils and a single β-sheet formed by strands β1, β2, and β8 (Gottipati et al., 2013). The first two β- strands of the sheet are formed by the first 100 amino acid residues in the sequence while the last

6 residues at the C-terminus (163–168) construct the final β-strand which folds back into the protease active site, positioning the C-terminus, Cys168, for cleavage and remains within the

substrate-binding pocket after cleavage. This explains why Npro cleaves only at its C-terminal end and is proteolytically inactive thereafter, as the active site is no longer accessible to cleave any other substrate molecules. The zinc-binding domain of Npro is produced by amino acid residues 101 through 156 and forms an anti-parallel β-sheet consisting of five β-strands, β3, β4,

β5, β6, and β7 (Zogg et al., 2013).

Site-directed mutagenesis studies resulted in detection of three essential residues for catalytic activity in Npro; Glu22, His49, and Cys69 (Rumenapf, et al. 1998; Rawlings et al.,

2012). However, detailed analysis of this protease domain revealed that these residues do not form a catalytic triad commonly found in many viral proteases. To this end, the position of

Glu22 was found to be too far from the active site to be involved in protease activity, instead,

Glu22 was found to form a salt bridge with Arg100 which imparts stability to the overall structure of Npro which explains the deleterious effect that Glu22 mutations have on catalytic activity of Npro (Rumenapf, et al., 1998). In contrast, the spatial arrangement of Cys69 and

His49 was determined to be sufficient to form a catalytic dyad responsible for protease activity of Npro. Cys69 was determined to be the nucleophilic residue in the active site of Npro, consequently, Npro is considered to be a cysteine protease. However, because Npro does not share sequence homology with any other known proteases, it has been assigned to its own family of cysteine proteases designated C53 (Rawlings et al., 2012).

Interestingly, it has been shown that the expression of Npro is not required for viral replication of CSFV in vitro (Tratshin et al., 1998). Additionally, the deletion of Npro from the genome of CSFV produces attenuation indicating that Npro may be a virulence factor of CSFV

(Mayer et al., 2004). To this end, Npro has been shown to be able to block the host cellular type I interferon responses to virus infection such as destruction of viral RNA, inhibition of cellular transcription and translation, and ultimately apoptosis. The central players of the signaling cascade that culminates in the induction of type I IFN are interferon regulatory factor 3 (IRF3) and IRF7. It has been demonstrated that Npro hinders the early innate immune response by binding IRF3 and targeting it for proteasomal degradation in the host cell (Bauhofer et al., 2007).

Concurrently, Npro has also been shown to be able to modulate the IRF7 turnover, limiting the

IFN-α induction in plasmacytoid dendritic cells (pDC) providing the virus with another strategy to elude the host innate defenses (Fiebach et al., 2011). Importantly, Npro relies on an intact Zn- binding domain for the interaction with both IRF3 and IRF7 (Szymanski M. R., et al., 2009;

Fiebach et al., 2011)

Core

The core (capsid) protein is a small, highly basic protein composed of 86 to 89 amino acids. The mature core protein of CSFV requires processing at the N and C termini in order to be released from the polyprotein. Processing occurs by three enzymatic cleavage events before incorporation into infectious virions. As described previously, the amino terminus of core is generated by autoprotease activity of Npro (Stark et al., 1993). The subsequent cleavage event is directed by a signal sequence in the carboxy terminus of core that targets the nascent protein to the endoplasmic reticulum (ER) membrane. Host cellular endoplasmic reticulum-associated signal peptidase (SPases) cleaves between Ala267/Asp268 in the ER lumen liberating Core protein from downstream Erns (Rumenapf et al., 1993). The final processing event is performed by host

signal peptide peptidase (SPPases) within the membrane spanning anchor of Core with the final

C terminus determined to be alanine 255 in CSFV (Heimann et al., 2006). The liberated mature

Core protein does not contain pronounced secondary structure and is primarily disordered

(Murray et al., 2008). Disordered, unfolded regions of viral capsid proteins are often involved in nucleic acid binding, as such, it has been shown that pestivirus Core protein binds to RNA, although with low affinity and little discernible specificity (Murray et al., 2008). It is likely that this relatively low binding affinity allows the RNA genome to be released and accessed for translation upon entry into a new host cell. Core proteins of all Flaviviridae exhibit RNA chaperone activity which was determined to be mediated by disordered, highly charged protein segments of Core (Ivanyi-Nagy et al., 2008). This RNA chaperone activity likely results in the condensation of the viral RNA genome and subsequent packaging into virions. Through mechanisms that are not well understood, the core protein and RNA genome together form the nucleocapsid and become enveloped by an ER-derived lipid membrane containing viral glycoproteins Erns, E1, and E2 (Thiel et al., 1991).

Experimental evidence based on the results of various deletion and truncations demonstrated that CSFV Core protein is indispensable for infectious virion formation (Reidel et al., 2010). However, more recently it was found that single amino acid substitutions within the

NS3 helicase domain are able to compensate for functionally compromised Core proteins even when nearly 90% of the core gene is deleted (Reidel et al., 2012).

In an effort to elucidate the biological function of CSFV Core, Gladue et al. utilized a yeast two-hybrid (YTH) approach to identify swine proteins that interact with Core protein of

CSFV strain Brescia. Screening revealed several host proteins that specifically bound to Core.

The most notable of these proteins included the cellular proteins SUMO-1 (small ubiquitin-like modifier), UBC9 (E2 SUMO conjugating protein), IQGAP1 (IQ motif containing GTPase activating protein1), and OS9 (osteosarcoma amplified 9 protein) (Gladue, et al., 2010; Gladue, et al., 2011; Gladue, et al., 2014). In general, the data obtained by characterizing each of these viral – host protein interactions suggests that the Core protein plays an important role in virus virulence.

The first two host cell proteins examined after YTH assay findings were, SUMO-1 and

UBC9, both of which are involved with the cellular SUMOylation pathway. It has been demonstrated, that the conjugation of SUMO can induce protein activation, affect protein stability, and cause changes in protein intracellular localization (Kerscher, et al., 2007).

SUMOylation has also been shown to play a role in the regulation of gene expression and maintenance of genomic stability (Seeler, et al., 2003). Several viruses, including human cytomegalovirus, Epstein–Barr virus, dengue virus, Moloney murine leukemia virus, and vaccinia virus, induce modifications to the host SUMOylation pathway in order to evade the host immune response (Sadanari et al., 2005; Huh et al., 2008; Shirai et al., 2008; Chiu et al., 2007;

Yueh et al., 2006; Palacios et al., 2005). The specific role of SUMOylation in CSFV was determined by initially mapping putative SUMOylation binding sites to two distinct regions in

CSFV Core protein. (Gladue, et al. 2010). Disruption of protein - protein interaction by substitution of specific residues within these regions was found to attenuate the virus in vivo.

Similarly, interactions of viral proteins with the SUMOylation pathway in other viruses has also been observed to effect viral infectivity by either preventing or inducing SUMO conjugation of

target proteins, as shown for Ebola Virus Zaire VP35, Adenovirus CELO Gam1, Human

Herpesvirus 6 IE2, and Human Cytomegalovirus IE2 (Chang et al., 2009; Chiocca et al., 2007;

Tomoiu et al., 2006 Ahn et al., 2001). Although the function of SUMOylation within CSFV

Core is still not clear, findings suggests that the interaction of Core with the cellular

SUMOylation pathway plays a significant role in the CSFV growth cycle as well as viral propagation in vivo (Gladue et al., 2010).

Subsequently, interaction of cellular protein, IQGAP1 with CSFV core was further examined to determine its significance. IQGAP1 protein has been shown to play an important role as a regulator of the actin cytoskeleton (Fukata et al., 2002; Watanabe et al., 2004). Notably, it has been reported that interaction of IQGAP1 with the Moloney murine leukemia virus

(MMLV) matrix (M) protein plays an important role in the intracellular trafficking of MMLV, which is essential for virus replication (Leung et al., 2006). In this same study the critical residues within MMLV M protein that interact with IQGAP1 were mapped by mutagenesis experiments. Based upon these findings, four regions within CSFV Core, designated I – IV, were identified that potentially bind IQGAP1. (Gladue et al., 2011). Amino acid residues in these putative binding sites were replaced with alanines. Resulting mutations in areas I and III completely disrupted the Core–IQGAP1 protein–protein interaction while substitutions in areas

bind cellular IQGAP1 protein during infection plays a critical role in virus virulence within the swine host.

Lastly, osteosarcoma amplified 9 protein (OS9) a protein involved in the endoplasmic reticulum-associated degradation (ERAD) pathway, was identified by YTH system to interact with CSFV Core (Gladue et al., 2014a). OS9 has previously been shown to be involved in the degradation of misfolded glycoproteins (Huttner et al., 2012), which could be significant considering CSFV Erns, E1 and E2 are heavily glycosylated structural proteins. Therefore, it is possible that binding of OS9 by the Core protein could prevent degradation of viral glycoproteins. Utilizing alanine scanning mutagenesis, the OS9 binding site within the CSFV

Core protein was mapped to the C-terminus of unprocessed Core, before its cleavage by signal peptide peptidase. A recombinant CSFV mutant lacking OS9 binding site in Core protein presented a significant decreased ability to replicate in primary swine macrophage cell cultures, the natural cell target of CSFV. However, no differences were observed in mutant infected swine when compared to animals infected with parental CSFV. As a result, it was concluded that the

OS9-Core interaction affects the efficiency of virus replication by an unclear mechanism. It is also possible that this interaction is a redundant virus function, as there was no observable decrease in virus virulence in swine. It appears that either the structural glycoproteins of CSFV are not recognized as misfolded proteins, or that the virus has multiple ways of preventing its glycoproteins from being degraded. Although the role for OS9 binding to the Core protein is not clear, it is clear that the OS9 binding site resides in the unprocessed Core protein. The role of unprocessed Core protein has not been previously reported in pestiviruses, however, it is possible that Core protein may function differently in both an unprocessed form and after cleavage of the

C-terminus. Overall, the YTH system and resulting identification of Core host proteins partners has provided interesting insight into the pathogenesis of CSFV by elucidating what appears to be multiple roles for Core in the CSFV replication cycle.

Erns Envelope Glycoprotein

Directly following the core protein is the first of three CSFV envelope glycoproteins,

Erns. This protein is specific to pestiviruses, with no corresponding homologue observed in any other genus within the Flaviviridae family (Lindenbach et al., 2001). Erns is a 44 – 48 kDa protein with a length of 227 amino acids. It contains 8–9 conserved cysteines that form intra- and intermolecular disulfide bonds. Erns is highly glycosylated with more than 50% carbohydrates in its mature form (Branza-Nichita et al., 2004). Erns is most commonly found as disulfide-linked homodimers (Thiel et al., 1991) with its most carboxy-terminal cysteine, C171, forming a disulfide bond between the Erns monomers (Tews et al., 2009). Of the three structural glycoproteins, only the structure for Erns has been determined (Krey et al., 2012). The crystal structure of Erns revealed that it contains five α helices and seven β strands with both a concave and convex face and is stabilized by four intramolecular disulfide bonds. Sequence analysis of different pestiviruses determined that the majority of N-glycosylation sites are localized to the convex face which is reasonable considering glycosylation on the concave face would likely interfere with Erns dimerization as well as enzymatic activity. Erns lacks a transmembrane domain nonetheless it is able to anchor itself to the membrane by a long amphipathic helix at its

C-terminus (Fetzer, et al., 2005). In addition, a considerable amount of Erns is virion bound and is secreted from infected cells into the extracellular environment (Rumenapf, et al., 1993).

Several functions have been assigned to Erns. First, comparative sequence analysis and subsequent biochemical analysis established that Erns possesses RNase activity (Schneider et al.,

1993). It is probable that this RNase activity of Erns antagonizes host immune response by possibly mediating degradation of extracellular RNA components that might otherwise interact with host cell intracellular pattern recognition receptors (PRRs) thus circumventing the host immune response to viral infection (Schneider et al., 1993). In addition, two mutations in Erns at residues H297 and H346 have been identified that eliminate Erns RNase activity, resulting in attenuation of the parental CSFV strain (Meyers et al., 1999). Erns has also been proposed to mediate initial attachment to cells by binding to glycosaminoglycans (GAG) on the host cell surface, although it is probable that this particular binding is not sufficient for entry into the cell as observed for many other viruses which require additional, more specific cell surface interaction to facilitate penetration of the host cell (Hulst et al., 2000). Furthermore, as a viral surface protein, Erns is also able to elicit virus neutralizing antibodies (Weiland et al., 1992) and as such can block pestivirus infection in tissue culture (Hulst et al., 1997) as well as provide protection in infected animals against lethal CSFV challenge (Hulst et al., 1993). While the Erns is not necessary for attachment to the host cell as observed with non-Erns pseudotyped virus carrying E1 and E2 glycoproteins (Wang, et al., 2004), Erns is however not indispensable as it is required for viral propagation (Widjojoatmodjo, et al., 2000).

As mentioned previously, Erns is heavily glycosylated with N-linked glycans accounting for nearly half of its molecular weight. N-linked glycosylation is important not only for correct protein folding and assembly but also for receptor binding, mediation of membrane fusion and

penetration into host cells, directing virus morphogenesis at the budding site, and acting as antigens to which neutralizing antibodies are directed (Shi et al., 2004). The importance of glycosylation varies between proteins; some proteins are completely dependent on glycosylation, whereas others display no dependence at all. In addition, it is often found that some glycosylation sites are more important than others when individually evaluated (Helenius et al.,

2001). For CSFV Erns, seven potential sites for the attachment of N-linked oligosaccharides were identified. In order to determine the importance of these individual glycosylation sites on

CSFV viral infectivity and virulence in swine, seven CSFV glycosylation deletion mutants were constructed (Fernandez-Sainz, et al. 2008). It was found that only one of the seven Erns mutants, with Asn - Ala substitution at Erns amino acid residue 269, had an impaired growth rate with a

5- to 10-fold decrease in the final virus yield. When these seven deletion mutant viruses were assessed in animals, only the mutant virus containing mutation of N269A resulted in an attenuated phenotype with decreased virus replication and shedding in comparison to parental

CSFV strain. In addition, these animals were completely protected from lethal CSFV challenge as earlier as 3 days post-inoculation. Interestingly, the rest of the single glycosylation site mutant viruses were virulent in swine, demonstrating that glycosylation at position 269 is important for in vitro and in vivo infectivity.

For many enveloped viruses, the pattern of glycosylation on glycoproteins has been demonstrated to affect the number of accessible epitopes recognized by the host immune system as antigen (Fournillier et al., 2001). Therefore, to further examine the role of Erns glycosylation on the induction of humoral immune response and protection against lethal challenge, animals were immunized with either purified glycosylated or nonglycosylated CSFV Erns subunit protein

expressed in baculovirus (Gavrilov, et al., 2011). Pigs vaccinated with glycosylated Erns were found to be protected against CSFV surviving lethal challenge without presenting clinical signs of disease. Conversely, animals inoculated with nonglycosylated Erns protein and challenged with virulent CSFV developed symptoms indistinguishable from those observed in mock vaccinated control animals. Despite the observed protection in glycosylated Erns immunized animals, levels of Erns antibodies and virus neutralizing antibodies were either low or undetectable depending on the number of times these animals were immunized with glycosylated

Erns before challenge. Consequently, glycosylation of glycoproteins can also play a role in masking viral neutralizing epitopes (Back et al., 1993), as a result, the removal of glycan moieties has been shown to enhance sensitivity of neutralizing antibodies and increase immunogenicity of viral epitopes (Reitter et al., 1998; Wei et al., 2003; Ansari et al., 2006).

Therefore, in order to determine whether the removal of individual glycosylation sites in Erns would alter the antibody response against CSFV, animals were immunized with mutated glycosylated forms of Erns subunit protein (Gavrilov, et al., 2011). All animals survived lethal challenge and were protected against clinical disease with the exception of two groups of pigs immunized with Erns N3 and Erns N7. Contrary to what has been observed for other viruses, such as Porcine reproductive and respiratory syndrome virus (PRRSV), simian immunodeficiency virus (SIV) and hepatitis C virus (HCV) (Ansari et al., 2006; Reitter et al.,

1998; Helle et al., 2010), removal of single glycosylation sites from Erns did not result in enhancement of the antibody response in vaccinated pigs. From these results it is clear that immunogenicity of Erns is dependent upon glycosylation pattern for the proper conformation necessary for eliciting a protective immune response against CSFV, although, the exact mechanism mediating that protection remains unknown.

E1 envelope glycoprotein

E1 is the only pestivirus envelope glycoprotein for which no structure or function has been determined. E1 is a 25–33kDa protein with a transmembrane anchor. Liberation of E1 from the polypeptide has been shown to occur in ER by mutation and inhibitor studies and that the cleavage between Erns and E1 is mediated by host cellular SPase despite not having a transmembrane anchor which is a common earmark in SPase cleavage sites (Bintintan et al.,

2010). Likewise, cellular SPase is also responsible for processing E1at its carboxy terminus

(Rumenapf et al.,1993). E1 contains several cysteine residues, most of which are well conserved throughout pestiviruses (Fernandez-Sainz et al., 2014). It is probable that these cysteine form intramolecular disulfide bonds which stabilize the protein structure. Importantly, E1 and E2 form a heterodimer via a disulfide bridge located on the virion surface (Weiland et al., 1990). Since the pestivirus E1 glycoprotein is approximately half the size of glycoprotein E2, recent models predict that E1is very thin and long in order to span the distance between the viral envelope surface and host cell membrane after E2 interacts with the host cell receptor (El Omari et al.,

2013).

While little is known about the functionality of E1, it has been determined that infected animals do not develop neutralizing antibodies against E1. Correspondingly, it has proven problematic to raise antibodies against E1 protein even when heterologously expressed with either Erns or E2, both of which individually generate neutralizing antibodies against CSFV. As a result of the lack of available antibodies against E1, detection of E1 is very difficult; therefore,

E1 is typically analyzed in conjunction with one of the other two envelope glycoproteins.

Accordingly, it has been determined that E1-E2 heterodimers are necessary to mediate attachment and virus entry into susceptible cells (Ronecker et al., 2008; Wang et al., 2004).

Furthermore, in a study designed to reveal the mechanism responsible for viral fusion, no apparent fusion peptides were revealed in the crystallized E2 structure, as a result it has been proposed that E1 might contain the obscure fusion peptide, with a stretch of hydrophobic amino acids between residues 57–85 identified as a possible candidate (El Omari et al., 2013). Lastly, unlike Erns and E2 which are both able to provide protection against lethal CSFV challenge, E1 alone was found to not be sufficient to elicit a protective immune response indicating that E1 by itself is not immunogenic (Konig et al., 1995).

In efforts to identify virulence and host range determinants of CSFV, Risatti et al. used transposon linker insertion mutagenesis (TLIM) to mutate an infectious clone of the pathogenic strain Brescia, BIC (Risatti et al., 2005b). A resulting viral mutant, RB-C22v, with a 57 nucleotide insertion in carboxyl terminus of E1 glycoprotein at genomic position 2429 was found to be highly attenuated in pigs. RB-C22v infected animals exhibited decreased replication in tonsils, limited generalization of infection, and significant reduction of viral shedding. Notably, these animals survived lethal challenge of CSFV presenting no signs of clinical disease. These results indicate that this mutation in E1 glycoprotein attenuates the virus and consequently has been identified as a virulent factor of CSFV.

As mentioned previously, E2 glycoprotein forms heterodimers with E1 glycoprotein via disulfide bridges between cysteine residues. The interaction between these envelope proteins plays an important role in virus assembly, virion maturity, and viral infectivity. The E1 envelope

protein of CSFV contains six cysteine residues, however, until recently the contribution of E1 cysteines in formation of disulfide bridges with E2 had not been determined. Furthermore, the biological significance of the formation of E1-E2 heterodimers during infection and the role of

E1 cysteine residues in CSFV replication or infection of swine was also not known. In order to begin to understand the role of these residues in the formation of heterodimers and virulence in vivo, a panel of single and double recombinant mutant viruses was constructed by substituting cysteine residues with serine in E1 glycoprotein CSFV (Fernandez-Sainz et al., 2011).

Interestingly, individual substitutions of cysteine residues did not have any impact on the formation of E1-E2 heterodimers or virulence in swine. Surprising 11 out of 15 double mutant constructs did not produce viable virus. However, of the remaining double mutant viruses, one exhibited an attenuated phenotype in animals while the other 3 retained the virulent phenotype of parental virus in pigs. In addition, the attenuated mutant displayed a partial disruption of E1-E2 heterodimer formation in infected cells as well as decreased growth rate in primary swine macrophages. Despite its diminished growth in vitro, this mutant virus provided protection in swine challenged with virulent CSFV at 3 and 28 days post infection. The attenuated virus contains mutations at amino acid position 24 and 94 in E1 glycoprotein suggesting that these residues are critical in the proper formation of E1-E2 heterodimers and virulence in swine.

Lastly, as with Erns, the role of post translational glycosylation on the immunogenicity and protective efficacy in pigs was evaluated for E1 both in the context of recombinant viruses as well as purified glycosylated forms of E1 glycoprotein expressed in baculovirus (Fernandez-

Sainz et al., 2009; Gavrilov, et al., 2011). Three putative N-linked glycosylation sites were identified in E1 (N500, N513 and N594). To determine the role of these glycosylation sites on

viral infectivity and virulence in swine a panel of E1 mutants were constructed in which each of the potential N-linked glycosylation sites was mutated either individually from Asn – Ala or in various combinations (Fernandez-Sainz et al., 2009). Substitutions at all three putative sites proved deleterious on virus yield, while all single and double mutants produced viable progeny.

When assessed in animals, only one of the single site mutants (N594A) and one of the double mutants (N500A/N513A) were found to be attenuated. In addition, these two recombinant viruses efficiently protected animals from virulent challenge at 3 and 28 days post-infection. It was concluded from these findings, that residue N594 in El envelope protein is critical for virus viability in the context of two or more glycosylation site modifications and E1mutant viruses

N594A and N500A/N513A lack determinants necessary for CSFV virulence in swine. More recently, purified glycosylated and unglycosylated E1 glycoproteins expressed in baculvirus were developed and assessed in swine (Gavrilov, et al., 2011). No detectable neutralizing antibodies were raised against either form of E1 subunit protein in immunized animals. In addition, neither form of E1 glycoprotein was able to provide protection against lethal CSFV challenge in swine. These findings indicate that envelope protein E1 is not immunogenic and as a result does not play a major role in immunity against CSFV.

E2 envelope glycoprotein

E2, the major envelope glycoprotein of the virus, has a molecular weight of 53–55kDa.

The cleavage between E2 and its adjacent proteins, E1 and p7, is processed by host cellular ER associated SPase (Elbers et al.,1996; Rumenapf et al., 1993). The C-terminus of E2 contains a

hydrophobic sequence that acts as transmembrane anchor. The mature E2 envelope protein is

373 amino acids long and contains 15 conserved cysteines which form intra-and inter molecular disulfide bridges (El Omari et al., 2013; Li et al., 2013). As described previously, E2 associates with E1 to form disulfide-linked heterodimers which are retained on the viral surface by their transmembrane anchors. In addition, E2 glycoprotein is also able to form E2-E2 disulfide-linked homodimers (Weiland et al., 1990). During virus assembly, E2 homodimers are formed early, whereas E1-E2 heterodimers are formed later after the release of E1 from the endoplasmic reticulum chaperone enzyme calnexin (Branza-Nichita et al., 2001).

The CSFV E2 glycoprotein exists as an elongated protein with four antigenic domains arranged in a linear fashion in the order of B/C/D/A (Wensvoort et al., 1989). All four domains are located in the N-terminal half of E2 ectodomain and form two individual antigenic units. The unit closest to the amino terminal portion of the E2 seqeunce is comprised of domains B/C while the other unit is composed of domains D/A (Wensvoort et al., 1989). The B/C domains are linked by a putative disulfide bond, whereas D/A domains are presumably bound by two putative disulfide bonds (van Rijn et al., 1994). Domains B/C and D/A each form a globular head connected by a pan-handle shaped link. Domain A has been subdivided further into A1, A2 and

A3 subdomains. A1 and A2 are conserved among classical swine fever virus strains, and domains A1, B and C are involved in neutralization (Wensvoort, et al., 1989). In contrast, subdomain A3 and domain D, are neither neutralizing nor conserved (van Rijn et al., 1993).

Functionally, E2 has been suggested to be responsible for receptor-binding and identified as the main target for neutralizing antibodies (Weiland et al., 1990). The E2 envelope protein has also been observed to play a major role in species tropism of pestiviruses (Liang et al., 2003;

Reimann et al. 2004). Importantly, both CSFV and BVDV use the same unknown host cell receptor as demonstrated by effective inhibition with either virus on porcine and bovine cells

(Hulst et al., 1997). In addition, the availability of a large panel of monoclonal antibodies against

E2 have made characterization of E2 possible by mapping binding sites and antigenic regions in order to gain information on the structure (Wensvoort G., 1986; van Rijn et al., 1993; van Rijn et al., 1994). Glycoprotein E2 is the most immunogenic of all CSFV glycoproteins (Konig et al.,

1995; Weiland et al., 1990). Moreover, it is well established that E2 alone is sufficient to elicit a protective immune response in animals against CSFV (Rumenapf, et al., 1991; van Zijl, et al.,

1991; Hulst et al., 1993; Hammond, et al., 2000). Consequently, due to these E2 glycoprotein specific properties, E2 has often been utilized in the development of potential CSFV vaccine candidates (van Gennip et al., 2000; Reimann, et al., 2004; Koenig et al., 2007).

Unlike E1 glycoprotein, the envelope glycoprotein E2 has been extensively studied and characterized. Accordingly, multiple regions within E2 have been identified as virulent factors of

CSFV (Risatti, et al., 2005a; Risatti et al., 2006). The role of E2 in virulence was first established in a study designed to identify genetic determinants of CSFV virulence and host range, in which a series of chimeric viruses were constructed where regions of CSFV genome were systematically replaced with the homologous regions of attenuated CSFV strain CS in the genetic backbone of highly virulent CSFV Brescia strain (Risatti, et al., 2005a). The resulting chimeric viruses were evaluated in swine for viral virulence. Interestingly, only chimeras harboring the CS E2 gene were attenuated suggesting that a significant virulence determinant absent in CS E2 was responsible for attenuation. Furthermore the exact area in CS E2 producing the attenuation was mapped between amino acid residues 886 and 1059 (Risatti, et al., 2005a, and 2007). These results demonstrate that CS E2 alone is sufficient for attenuating Brescia,

indicating that CSFV E2 glycoprotein is a major determinant of virulence in swine. In a subsequent study, progressive mutations were introduced into an area of E2 between residues

829 and 837 (Risatti, et al., 2006) that contains a distinct CSFV specific epitope recognized by monoclonal antibody (mAb) designated WH303, a reagent which fails to react with BVDV or

BDV E2 and is routinely used for CSF diagnostics (Lin et al., 2000). Amino acids in mAb

WH303 epitope, TAVSPTTLR, of CSFV Brescia were progressively replaced with the corresponding amino acid residues of BVDV, TSFNMDTLR, producing 4 chimeric viruses. The resulting viruses were designated T1v – T4v and were evaluated in animals. T1v – T3v induced lethal disease, while T4v induced mild and transient disease in pigs. Notably, animals infected with T4v were protected when challenged with virulent Brescia virus at 3and 21 days post- vaccination. These results demonstrate residues, TAVSPT, in mAb WH303 eptiope of E2 play a significant role in CSFV virulence.

The majority of viral integral membrane proteins are post translationally modified by addition of N linked carbohydrate chains, although some have only O-linked carbohydrate chains while others have both (Doms, et al., 1993). Accordingly, five putative N-linked sites and one putative O-linked glycosylation have been predicted by algorithm analysis for the E2 glycoprotein (Moormann, et al., 1990). Notably, a sixth N-linked putative glycosylation site has been reported for several CSFV strains including CSFV Brescia strain. Importantly, predicted E2 glycosylation sites are highly conserved among CSFV isolates. However, not all predicted sites in a protein sequence are used for carbohydrates, since many of them are either inefficiently glycosylated or remain unglycosylated (Shakin-Eshleman, et al., 1992; Gavel, et al., 1990). As such, Risatti et al. investigated the role of E2 glycosylation in the highly virulent CSFV Brescia

strain in swine (Risatti et al., 2007). Seven putative glycosylation sites in E2 were modified by site-directed mutagenesis of a CSFV Brescia infectious clone (BIC) creating a panel of 15 mutant viruses used to investigate whether the removal of putative glycosylation sites in the E2 glycoprotein would affect viral virulence and pathogenesis in pigs. Interestingly, no viable virus was produced when all putative glycosylation sites were removed; however, viable progeny were generated upon reversion at mutation N185A back to wild type Asp. Furthermore, this reversion mutant virus was found to be attenuated in swine. In addition, analysis of single E2 glycosylation site mutant viruses in pigs revealed that removal of glycosylation at amino acid N116A (N1v virus) was responsible for BICv attenuation. N1v efficiently protected swine from challenge with virulent BICv at 3 and 28 days post-infection, suggesting that glycosylation at amino acids N116 and N185 are critical for E2 activity as well as viral virulence. In addition, the importance of glycosylation in the immunogenicity of E2 was assessed using purified glycosylated, or nonglycosylated versions of E2 expressed in baculovirus. (Gavrilov, et al., 2011). Pigs vaccinated with glycosylated E2 protein induced virus neutralizing antibodies and were protected against lethal challenge with CSFV. Conversely, animals inoculated with nonglycosylated E2 subunit protein did not elicit detectable neutralizing antibodies and when challenged with virulent CSFV developed symptoms indistinguishable from those observed in mock vaccinated control animals (Gavrilov, et al., 2011). Subsequently, naïve animals were immunized with single site glycosylation mutated E2 subunit proteins to assess whether individual glycosylation sites present in E2 were critical in the induction of the neutralizing antibody response.

Neutralizing antibodies were detected in sera from all pigs immunized with single site glycosylation mutant E2 proteins. While, most of the animals infected with E2 single glycosylation mutant viruses presented diminished levels of neutralizing antibodies in

comparison to those animals vaccinated with glycosylated E2 subunit protein, two groups of animals infected with single mutations at either amino acid residue 260 or 297 elicited neutralizing antibody levels indistinguishable from levels raised in animals immunized with the glycosylated E2 subunit protein. In addition, all animals infected with single site glycosylation mutated forms of E2 protein were protected against lethal CSFV challenge, except for animals infected with E2 glycosylation mutant protein in which the glycan moiety at position 121 was removed. Overall, these results demonstrate that glycosylation pattern plays a major role in the immunogenicity of CSFV E2 glycoprotein

As mentioned previously, E2 glycoprotein is involved in cellular attachment and entry into susceptible host cells. Entry of enveloped animal viruses requires fusion between the viral membrane and a cellular membrane. Using proteomic computational analysis, a putative fusion peptide (FP) between amino acid residues 818 to 828 was identified in CSFV E2. (Garry et al.,

2003). This sequence, CPIGWTGVIEC, was found to be highly conserved among pestiviruses, as is true of fusion peptides from other enveloped RNA viruses. Fernandez-Sainz et al. (2014) evaluated the role of this putative fusion by generating a series of 8 recombinant cDNA constructs by mutating amino acids in the E2 FP region either individually or combinatorially in the genetic background of full-length BIC. Six of these constructs produced viable viruses while a double mutant, 818S/C828S, and triple mutant, P819S/I820S/W822S amino acid substitutions were deleterious for CSFV replication. Amino acid substitutions of the remaining 6 FP mutant viruses were not found to significantly affect either virus growth in vitro or virulence in animals.

Notably, the lethal amino acid substitutions of the double and triple mutants did not appear to impact the secondary structure of the FP at all. In contrast, the triple mutation was shown to

impair the ability of a synthetic peptide harboring these amino acid substitutions to penetrate target membranes. In addition, it was demonstrated that the synthetic peptide harboring the WT

FP sequence was more efficient at pertubing the lipid bilayer than synthetic peptides containing the double or triple amino acid substitutions. Overall, this preliminary characterization suggests that the E2 putative FP is involved in membrane fusion activity and plays a critical role in virus replication.

Lastly, until recently there was no information regarding host binding partners for the E2 glycoproteins which could help expand the current understanding of the functions of the E2 protein. To this end, Gladue et al. utilized the yeast two-hybrid system and identified 57 host proteins that bind the E2 glycoprotein of CSFV (Gladue et al., 2013). In an effort to map binding sites on E2 with these cellular proteins an alanine scanning approach was utilized thereby generating a panel of 76 CSFV E2 mutant proteins. However, after testing of all 76 E2 mutants against all the host proteins, identification of specific binding sites was unsuccessful as all of the host proteins were capable of binding all of the E2 alanine mutants. These results, suggest that the host proteins do not bind to linear amino acid stretches within of E2 rather it is likely that most of these proteins bind to multiple non-linearly arranged residues that are not disrupted by the polyalanine mutations in linear amino acid stretches. While no specific binding sites in E2 were identified, the possible roles for viral E2 interaction with these host proteins were speculated based upon analysis with the PANTHER classification system (Protein ANalysis

THrough Evolutionary Relationships). Fifteen different biological processes were found for these host proteins which include metabolic, cellular, developmental, immune system, cell communication, cell adhesion, system processses, response to stimuli, cellular compartment

organization, cell cycle, transport, generation of precursor metabolites and energy, apoptosis, localization, and reproduction processes.

Non structural protein p7

Immediately downstream of E2 is the gene coding for protein p7, a small hydrophobic polypeptide with a molecular weight of 6 to 7 kDa. This protein is not detected in virions of pestiviruses and is therefore classified as a non structrural protein (NSP) (Elbers et al., 1996).

Similar to many CSFV proteins described previously, p7 is released from the polypeptide by cellular SPase (Elbers et al., 1996). Notably, in pestivirus-infected cells, p7 has been detected as both an individual protein as well as part of fusion protein E2-p7 (Elbers et al., 1996). The existence of fusion protein E2-p7 is unusual since cleavage by SPase is extremely efficient. No precursor-product relationship was found between E2-p7 and E2, which suggests E2-p7 fusion protein must be functionally important (Harada, et al., 2000). Furthermore, the cleavage of precursor E2p7NS2 into mature p7 and E2-p7 has been found to be conserved among pestiviruses as well as hepatitis C virus (HCV). Therefore, it has been proposed that E2-p7 fusion protein may play an important role in the viral life cycle (Harada et al., 2000). However, upon further investigation it was found that the E2-p7 fusion protein is dispensable for both viral

RNA replication and generation of infectious virions. Consequently, the function of E2-p7 fusion protein is presently unknown. Conversely, it has been demonstrated that p7 is required for generation of infectious virions in pestiviruses (Harada et al. 2000; Gladue et al., 2012). Given that p7 contains two hydrophobic stretches of amino acids interrupted by a short charged segment which is predicted to form two transmembrane helices connected by a charged cytosolic

loop, it has been suggested that CSFV p7 may be a member of the viroporin family (Harada, et al., 2000). Viroporins are small polypeptides that are capable of modulating membrane permeability and may play a crucial role in the release of virions. Recently, the p7 protein of

CSFV was confirmed to function as a viroporin with pore forming activity by analysis in a liposome model membrane system mimicking the lipid composition of the ER membrane

(Gladue et al., 2012). Importantly, the pore forming activity was mapped to the carboxyl half of the p7 protein. In the same study, constructs containing in-frame partial deletions of p7 in the context of full length cDNA CSFV infectious clone were unable to generate infectious particles suggesting that p7 is essential for in vitro viral replication. In addition by an alanine mutagenesis screening approach, p7 mutant viruses were produced and tested in swine to determine the effect that alterations of different regions of p7 have on virulence. Some of the constructs where unable to produce infectious progeny while other constructs produced viruses that were either partially or completely attenuated in animals, indicating that p7 plays a significant role in virulence. More recently, Largo et al. (2014) mapped the minimal p7 sequence required for pore formation to amino acids 33–67 which encompasses the basic cytosolic loop as well as the c-terminal transmembrane in CSFV (Largo et al., 2014). While the c-terminal transmembrane was sufficient for pore formation, the n-terminal loop was shown to be necessary for regulating the process. Other target membranes were also examined in this study and it was found that the c- terminal hydrophobic domain was capable of inserting into membranes of the secretory pathway such as the ER and Golgi but not the plasma membrane.

NS2

NS2 of pestiviruses consists of about 450 amino acids. The N-terminal half of the protein is highly hydrophobic and may form up to seven transmembrane regions followed by a cytosolic domain. The cytosolic domain contains a putative Zn2+-binding site, consisting of four cysteine residues, which is an integral part of an autoprotease subdomain responsible for cleaving NS2-3 in cis between NS2 and NS3 (de Moerlooze et al.,1990; Lackner et al., 2004). After cis- cleavage, the C-terminus of NS2 stays in the active site of the protease, thereby preventing further NS2 cleavages in trans (Lackner et al., 2006). In addition, it has been found that proteolytic activity of NS2 depends on interaction with a cellular chaperone Jiv (J-domain protein interacting with viral protein) (Lackner et al., 2006). Notably, cleavage of NS2-3 is limited by the availabity of cellular Jiv as the complex formation between Jiv and NS2 is tightly regulated with one molecule of Jiv responsible for the release of one NS3 protein.

Recently, several amino acid residues were identified in the N-terminal region of NS2 that are critical in regulation of the CSFV life cycle (Li et al., 2015). In this study, residues at position 60 and 78 in the NS2 protein were found to be independently essential for RNA replication and virus production. While a substitution at amino acid residue 100 in NS2 resulted in significantly reduced viral titers suggesting that NS2 may also play a role in regulation of

CSFV secretion from infected cells.

NS2-3

Interestingly in pestiviruses, the unprocessed NS2-3 precursor forms a stable protein in infected cells (Collett et al., 1991). As a result, NS2-3, NS2 and NS3 proteins are all present within infected cells at varying concentrations. It was once thought that the difference observed between pestivirus noncytopathogenic (noncp) and cytopathogenic (cp) biotypes were dependent upon the processing of NS2-3 (Thiel, et al., 1996). Essentially, processing of NS2-3 to NS2 and

NS3 was suggested to result in the accumulation of NS3 which was responsible for the cytopathic effect observed in tissue culture infected with cp pestivirus strains. Conversely, this

NS2-3 processing was presumed to be absent in the noncp biotypes (Kummerer, et al., 2000).

However, it has been demonstrated that NS3 is indispensible for pestiviral RNA replication regardless of the specific biotype and cannot be functionally replaced by NS2-3 (Lackner et al.,

2004). Nevertheless, the significance of uncleaved NS2-3 in infected cells was ultimately revealed as being required for production of virus particles. This role has been illustrated in both

BVDV and CSFV where the genomes were modified in such a way as to prevent expression of

NS2-3 while allowing normal expression of all the other viral proteins, including NS2 and NS3

(Agapov et al., 2004 ; Moulin et al., 2007). In both instances, the absence of uncleaved NS2-3 correlated with a defect in the production of infectious virus progeny, despite evidence of efficient viral RNA replication. Importantly as mentioned earlier, cleavage of NS2-3 is temporally modulated by the cellular cofactor Jiv which associates with NS2 (Lackner et al.,

2006). This NS2-mediated cleavage acts as a switch between RNA replication and infectious particle formation. Once the Jiv supply in the infected cell is consumed cleavage of NS2-3 ceases resulting in decreased viral RNA replication due to NS2-3’s inability to replace NS3 in the

replication complex (Lackner et al., 2006). Uncleaved NS2-3 accumulates in the cell and associates co-translationally with NS4A and subsequently enters the virion assembly process.

NS3

The NS3 protein is the most highly conserved non-structural protein among pestiviruses and predicted to be hydrophilic with a net positive charge (Xu et al., 1997). It has been established that NS3 plays a key role in formation of the replication-transcription complex and subsequent RNA replication (Zhang et al., 2003). NS3 is a multifunctional protein containing protease, helicase and NTPase activity. In the N-terminal portion of NS3 resides a chymotrypsin- like serine protease. This chymotrypsin-like serine protease acts in both cis-cleavage (NS3-4A) and trans-cleavage (NS4A-4B-5A-5B) (Rice et al., 1996). The other functional domains of NS3 are found in the C-terminal region encompassing the helicase and NTPase domains (Warrener et al., 1995). Notably, the NS3 serine protease requires the stable association with its cofactor

NS4A in order to reach full activity (Xu et al., 1997). Consequently, the NS3/4A protease is responsible for cis-cleavage generating the C-terminus of NS3 as well as all downstream cleavages which occur in trans, releasing NS4B, NS5A, and NS5B from the polyprotein

(Wiskerchen et al., 1991). Similarly, uncleaved NS2–3 was also shown to possess serine protease activity, though with reduced activity, and as a result could not functionally substitute for NS3 in viral RNA replication (Lackner et al., 2004). Once cleaved by NS2, the liberated NS3 protein interacts with NS4A as well as with the downstream nonstructural proteins of the replication complex at the membrane of the endoplasmic reticulum where negative-sense RNA templates and progeny viral genomes are produced (Xu et al., 1997). For these processes, the

helicase and NTPase activities of NS3 were also found to be essential (Gu et al., 2000). Using mutational analysis, Gu et al. (2000) demonstrated that the NS3 NTPase/helicase activities of the

BVDV were essential for the synthesis of minus-strand RNA. In addition, they also found that

NTPase activity is absolutely required for helicase activity. While the functional mechanism of helicase activity remains unclear it has been proposed that NTPase/helicase could allow initiation of minus-strand template synthesis by unwinding the secondary structures present in the 3’ UTR thereby allowing initiation by the RNA-dependent RNA polymerase (RdRp).

Lastly, a new and unexpected role has recently been described for the C-terminal region of NS3 of CSFV. In an experiment investigating properties of the pestivirus core protein C, a pestivirus mutant virus lacking almost the entire C gene sequence was unexpectantly found to be viable as a result of a random mutation in the helicase domain of NS3 (Riedel et al., 2010).

Further investigation demonstrated that single amino acid substitutions in the helicase domain of

NS3 could sufficiently compensate for the loss of core protein (Riedel et al., 2012).

Correspondingly, this role of the NS3 helicase domain has also been observed in virion morphogenesis in other members of the Flaviviridae family, such as yellow fever virus and HCV

(Patkar et al. 2008; Ma et al., 2008). Riedel et al. (2012) propose that the ability of NS3 to functionally replace the core protein may suggest that NS3 had a packing function in an ancestral prototype virus in the absence of a capsid gene which was later acquired during the evolutionary process in order to infect a novel host (Riedel et al., 2012).

NS4A

NS4A is a non-structural protein with a predicted weight of 10 kDa. It contains a hydrophobic, membrane spanning N-terminal region and a C-terminal cytosolic domain (Liang et al., 2009). NS4A has a length of 64 amino acid residues and is quite acidic. It is also highly conserved among pestiviruses. Early studies demonstrated that the BVDV NS3 serine proteinase expressed alone was not sufficient for cleavage at the 5A/5B site (Wiskerchen et al., 1991). It was later determined that NS4A forms a complex with NS3 serine proteinase domain acting as a cofactor for NS3 and together are essential for cleavage at the 4B/5A and 5A/5B sites (Xu et al.,

1997). In addition to this protease cofactor function, trans-complementation experiments revealed another critical role for NS4A in virion morphogenesis (Agapov et al., 2004; Moulin et al., 2007). Consequently, it was ascertained for both BVDV and CSFV that uncleaved NS2-3 requires expression of NS4A for optimal production of infectious virions (Agapov et al., 2004;

Moulin et al., 2007). Furthermore, Moulin et al. (2007) demonstrated that CSFV infectious particle rescue was dependent upon a combination of uncleaved NS2-3 and processed NS4A rather than the NS2-3-4A polyprotein precursor. Notably, the role of NS4A with NS2-3 in infectious particle formation is independent of NS4A function in conjunction with NS3 in RNA viral replication.

NS4B

NS4B is 347 residues long with a molecular mass of approximately 35kDa (Xu et al.

1997). It is a hydrophobic protein that is indispensable in pestiviral RNA replication (Collett et

al., 1991; Grassmann et al., 2001). NS4B is an integral membrane protein associated with the endoplasmic reticulum that possesses four putative transmembrane domains flanked by cytoplasmic N- and C-terminal regions (Weiskircher et al., 2009). The N-terminal region contains a conserved amphipathic helix that can become a transmembrane domain upon NS4B cleavage (Elazar et al., 2004) while the C-terminal region contains two predicted helical domains

(Jones et al., 2009). A recent study mapped amino acids in the N-terminal region of NS4B that are essential in the formation of a functional replication complex (Gouttenoire et al., 2009). In

HCV and BVDV replication studies, it has been demonstrate that NS4B colocalizes with replicase proteins NS3, NS5A and NS5B which suggests that NS4B may be a component of the

BVDV replication complex (Piccininni et al., 2002; Weiskircher et al., 2009). In addition, it has been shown that NS4B may also play an important role in virus assembly and release (Jones et al., 2009). Finally, it has been reported for multiple flaviviruses that NS4B is able to strongly block the IFN-induced signal-transduction cascade by interfering with STAT1 function thereby evading the host immune system (Muñoz-Jordan, J. L. et al., 2003 and Tasaka, et al., 2007).

Amino acid sequence analysis of pestivirus NS4B proteins revealed sequences with similarity to nucleotide-binding motifs (NBM), termed Walker A and B, which were initially observed in HCV although the location of these motifs within the protein differ among genus

(Gladue et al., 2011; Einav et al. 2004). In pestiviruses, the motifs’ sequences are located towards the C-terminal portion of NS4B, consisting of amino acid residues 209-216 and 335-342 of the protein, for Walker A and B respectively. This structural motif has been shown to bind and hydrolyze ATP, and GTP (Einav et al., 2004; Thompson et al., 2009). Substitutions of critical residues within these motifs have been reported to drastically reduce the ATPase and GTPase

activities of bacterial expressed NS4B proteins (Gladue et al., 2011). Similar mutations introduced into the genetic background of a full-length cDNA copy of CSFV strain Brescia rendered no infectious progeny or viruses with diminished replication capabilities, indicating that this NTPase activity is essential for the CSFV replication cycle (Gladue et al., 2011).

In addition to the Walker A and B motifs, a putative Toll/interleukin-1 receptor (TIR)- like domain has been predicted to reside in NS4B protein. This domain is unique to both CSFV and BVDV and is not found in other pestiviruses or flaviviruses. TIR domains participate in signal transduction of Toll-like receptors (TLRs). In general, TLRs are a large family of proteins that play a crucial role in recognition of various microbial components, activate the innate immune system, and induce of cytokines needed for the adaptive immune response (Akira et al.,

2004). Importantly, the TIR domain contains 3 conserved motifs, termed box 1, 2 and 3. It has been shown that alanine substitution in box 1 and 2 of a TIR domain in a human interleukin receptor can inhibit its signaling, while signaling was unaffected by alanine substitutions in box3 of the TIR domain. Based on these observations, Fernandez-Sainz et al. (2010) selectively generated mutations in boxes 1 and 2 in order to elucidate the function of this TIR domain in

CSFV NS4B. Mutations, V2566A, G2567A, and I2568A in the box 2 domain produced a virus

(NS4B VGIv) with an attenuated phenotype in pigs. Infection with recombinant NS4B VGI was characterized by localized replication in the tonsils with limited viral dissemination during infection. Furthermore, animals inoculated with NS4B VGIv were completely protected against lethal CSFV challenge as early as 3 days post infection. Analysis of cellular gene expression of infected primary swine macrophage revealed cultures infected with NS4B VGIv showed increased activation of several genes, compared to those in cultures infected with the parental

virus. Notably, most of the genes exhibiting increased transcriptional activation with NS4B

VGIv infection are similarly induced by activated TLR-7 (Lee et al., 2003). Furthermore, TLR-7 functional involvement was indirectly confirmed by the observation that transfection of macrophages with wild-type NS4B reduces the responsiveness of TLR-7 to a well characterized activator, imiquimod R837 (I-R837) while transfection with NS4B gene harboring box 2 domain mutations was less efficient at mediating this reduction. Ferandez-Sainz et al. (2010) speculated that these findings suggest that NS4B of wild-type CSFV blocks TLR-7 transcriptional activation during infection in animals.

NS5A

NS5A protein is a large, hydrophilic phosphoprotein with an approximate molecular mass of 58 kDa and is comprised of 497 amino acids. It encompasses three domains interconnected by two low-complexity sequences (LCSs). At the N-terminus of NS5A is an amphipathic α-helix which serves to anchor the NS5A protein to intracellular membranes (Brass et al., 2002). It was demonstrated that the first 27 to 33 amino acids of NS5A of hepaciviruses and pestiviruses are necessary and sufficient for both targeting and anchoring the NS5A protein to the membrane

(Brass et al., 2007). In addition, a conserved zinc-binding motif was also found in domain I of the NS5A protein of hepaciviruses and pestiviruses (Tellinghuisen et al., 2004; Tellinghuisen et al., 2006). Sequence analysis, identified four conserved CSFV cysteine residues, C2717-C2740-

C2742-C2767, involved in zinc binding (Sheng et al., 2010). Furthermore, mutation at any of these cysteine residues results in an NS5A protein unable to bind zinc a function shown to be

indispensable for RNA replication (Sheng et al., 2010). In addition, HCV and related viruses seem to be phosphorylated by the similar cellular kinase(s) which suggests that phosphorylation of the NS5A/NS5 proteins or their association with cellular kinases may play an important role in the flavivirus life cycle. While NS5A function is still not entirely clear, it is believed to be a component of the replication complex (Brass et al., 2002). NMR structure analyses suggests that the N terminus amphipathic helix may constitute a unique platform essential for the formation of the replication complex (Sapay et al., 2006). Finally, NS5A has recently been shown to regulate viral RNA synthesis and viral replication by either binding to the 3’UTR of the viral RNA genome or by modulation of NS5B RdRp activity by direct protein–protein interactions (Sheng et al., 2012).

NS5B

The last protein in the carboxyl end of the pestiviral polyprotein is NS5B which has a weight of about 77 kDa and contains 718aa. The CSFV NS5B protein has been demonstrated to contain RNA-dependent RNA polymerase (RdRp) activity and is able to bind to the 3′UTR in order to initiate genome replication (Steffen et al., 1999). RdRp is a required for replication of the positive-strand RNA genome. NS5B transcribes the genome into negative-sense strands, which serve as templates to produce more positive-sense RNAs for packaging into progeny viral capsids (Gong et al., 1996). The overall structure resembles a right hand with fingers, palm, and thumb domains and has 5 motifs designated A, B, C, D and E (O'Reilly et al., 1998). Motifs A,

B, C and D are located at the catalytic portion of the palm domain (Hansen et al., 1997).

Importantly, motif C forms a β-strand-turn-β-strand structure which contains the highly

conserved GDD motif found in all RdRps (Kamer & Argos, 1984). The GDD sequence is postulated to be involved in the catalytic-activity and metal-ion regulation of the RdRps

(Jablonski et al., 1991). It has been demonstrated that NS5B directly binds to NS3/4A (Ishido et al., 1998), NS5A (Yamashita et al., 1998), and indirectly to NS4B by help of NS5A (Lin et al.,

1997), as such NS5B is not only the central enzyme for the viral replication but may also be a core protein for assembly of the replication complex. Moreover, NS5B has recently been implicated to play a role in infectious virion assembly (Ansari et al., 2004).

Viral Life Cycle

The proposed replication strategy for pestiviruses and other members of the Flaviviridae family is far from being completely understood and is largely based upon studies done with flaviviruses. Consequently, the virus is believed to enter a susceptible host cell via a multistep process which involves cellular receptors specific for viral envelope glycoproteins (Lindenbach et al., 2001). It is presumed that the viral Erns glycoprotein initially interacts with a currently unknown host cell surface receptor which mediates attachment of the virion to the cell surface

(Gollins et al., 1985). Thereafter, the viral E2 glycoprotein is thought to interact with a separate and more specific cellular surface molecule facilitating penetration into the cell (Hulst et al.,

1997). It is speculated that the virion is then taken up into a prelysosomal endocytic compartment where the compartment’s acidic environment induces fusion between the viral envelope and the endosomal membrane thereby releasing the viral nucleocapsid into the cytosol of the host cell

(Lindenbach et al., 2001). Next, uncoating of the nucleocapsid by a process that is not yet fully understood releases the viral genome into the host cytoplasm where the viral genome due to its

positive polarity is immediately translated into a single polyprotein. Processing of the polyprotein into discrete proteins occurs co- and post-translationally by viral proteases and host cell proteases residing in the lumen of the endoplasmic reticulum (ER). Once the polyprotein processing is completed, the formation of RNA replicase from the nonstructural proteins leads to replication of the viral genome which takes place on the intracellular membranes (Xiao et al.,

2004). Viral assembly proceeds on the surface of the ER, where the structural proteins and the newly synthesized RNA bud into the lumen of the ER. As the immature viral particles pass through the secretory pathway, maturation of virions occur and the envelope proteins become glycosylated (Westaway, E. G., 1987). Finally, the nascent virions pinch off from the trans-Golgi network as exocytic vesicles which fuse with the plasma membrane releasing mature virions from the host cell (Mukhopadhyay et al., 2005).

Translation and Protein Processing

After the viral genome is released into the cytoplasm of the host cell the 5’UTR of the genome directs the RNA to ribosomes which are in close proximity with the ER (Leyseen et al.,

2000). The 5’UTR contains a complex RNA structure known as an internal ribosomal entry site

(IRES) which promotes the binding of the ribosome to the start codon thereby initiating translation of the viral RNA. Translation produces a single long precursor polyprotein which is processed into individual proteins co- and post-translationally.

The polyprotein processing begins with cleavage of Npro from the downstream capsid protein and is expected to occur co-translationally (Tratshin et al., 1998). The Npro, as stated

before, possesses autoproteolytic activity allowing it to cleave itself from capsid protein creating its own C- terminus and the N- terminus of capsid protein (Stark et al., 1993). Next, processing of the structural proteins (C, E0, E1, and E2) proceeds in three separate cleavage events, all of which are presumed to be catalyzed by host cell signalases residing in the lumen of the ER

(Rumenapf et al., 1993). These events lead to the formation of two intermediate products, E012 and E01, which are not present in the mature virion. The first step in this process is the translocation of an internal signal peptide, downstream of the capsid protein, across the ER membrane. Immediately, upon translocation of this signal peptide through the ER membrane, host signalase cleaves the capsid protein from the nascent E012 polyprotein (Rumenapf et al.,

1993). The transfer of E012 continues until the process is interrupted by a hydrophobic stretch of amino acids at the end of the E1 protein. This stretch of amino acids acts as a stop transfer signal and anchors the E1 protein to the membrane. Soon after, a second stretch of hydrophobic amino acids is encountered in the E012 polyprotein that acts as a signal sequence to begin the transfer of E2 through the ER membrane (Rumenapf et al., 1993). Translocation of E2 proceeds until interrupted by a set of stop/start transfer signals at the end of the E2 protein which acts as a second anchor tethering E2 to the membrane. The first intermediate precursor, E012, is formed upon the translocation of E2 at which point signalase rapidly cleaves E012 from the downstream translation product, p7. Processing of the E012 precursor continues with the cleavage of E2 resulting in the formation of the second intermediate precursor, E01 (Rumenapf et al., 1993). For reasons that remain unclear the cleavage between E1 and E2 occurs in a slightly delayed manner compared to the prior cellular signalase cleavage events. Finally, processing of the E01 precursor ensues soon after E2 is released thereby forming the two final glycoproteins, E0 (Erns) and E1.

E0 which does not appear to possess a membrane anchor presumably remains associated with the

budding virus through noncovalent associations with the other glycoproteins (Rumenapf et al.,

1993).

The next proteins in the polyprotein to be processed are the nonstructural proteins p7 and

NS2-3, both of which are largely composed of hydrophobic amino acids. It is believed that these proteins like the structural proteins are translocated across the ER membrane and processed into individual proteins by cleavages mediated by host signalase (Harada et al., 2000). Release of the remaining nonstructural proteins located downstream of NS2-3 is mediated by a serine protease within NS2-3 (Harada et al., 2000). This serine protease located near the 5’ end of the NS3 protein is essential for generation of its own C-terminus and for processing of the downstream cleavage sites (Tautz et al., 1997). Once NS4A is cleaved from the downstream product it acts as a cofactor to the NS3 serine protease for the remaining cleavages of NS4B, NS5A and NS5B

(Xu et al., 1997). Cleavage between NS2-3 and NS4A and cleavage between NS4B and NS5A occur very rapidly. Conversely, the cleavage between NS4A and NS4B and cleavage between

NS5A and NS5B are both delayed in comparison. The delay at these cleavage sites is thought to be dependent upon the composition of the amino acids surrounding the cleavage sites (Xu et al.,

1997).

RNA Replication

Viral replication of the genome is able to proceed only after processing of the polyprotein into individual proteins is complete because the nonstructural proteins NS4B, NS5A and NS5B are all required to form the viral RNA replicase. It is believed that NS5B encodes for RNA

dependent polymerase while the hydrophilic protein products of NS4B and NS5A are components of viral RNA replicase. Once the replication complex is assembled from the nonstructural proteins, it is able to recognize and bind to the 3’UTR of the viral genomic RNA and RNA synthesis is initiated (Xiao et al., 2004). RNA replication occurs in smooth membrane vesicles that appear as vesicle packets (VPs) (Lindenbach et al., 2001). These VPS are associated with the perinuclear membrane (Lindenbach et al., 2001). RNA replication begins with the synthesis of a negative-stranded RNA which serves as a template for the replication of additional positive-stranded genomic RNA (Lindenbach et al., 2001). RNA synthesis is asymmetric leading to 10 to 100 fold excess of positive strands over negative strands (Lindenbach et al., 2001). As described previously, the positive stranded progeny associate with nascent viral proteins for viral assembly and bud into the lumen of the ER, pass through the secretory pathway and are released as a mature virion into the extracellular space via exocytosis.

Clinical Signs of CSFV

CSFV causes a disease in swine with varying degrees of severity classified into three general forms: acute, chronic, and late onset. The course of disease depends predominantly upon the virulence of the viral strain and the age of the host animal but may also be affected by breed and immune status of the animal. Field isolates and laboratory strains of CSFV display a wide range of virulence and are distinguished as highly virulent, moderately virulent, low virulence and/or avirulent according to the severity of symptoms produced in swine. Highly virulent strains typically induce an acute phenotype of disease in infected animals with a 100% mortality rate, while moderate and low virulent strains generally induce a prolonged chronic form of disease

with mortality rates reaching as high as 90% (van Oirschot, J. T., 1986). Notably, it has been observed that the same virus isolate with moderate virulence can cause different forms of disease depending on the age of the infected animal, with younger animals commonly developing a more severe course of infection than older animals (Moennig et al., 2003). Not only do these differences in the degree of severity of infection make diagnosis of CSF difficult for veterinarians but in addition the symptoms of CSF are quite similar to those of many common farm infections such as salmonellosis, pasteurellosis, erysipelas, streptococcosis, and leptosspirosis which is further complicated by the emergence of new diseases such as porcine reproductive and respiratory syndrome (PRRS), porcine dermatitis and nephropathy syndrome which also have similar symptoms (Done et al., 2000).

The initial symptoms observed in both the acute and the chronic forms of CSF are depression, lethargy, anorexia, rhinitis, and conjunctivitis. Infected animals typically huddle together which is a visual indicator of the presence of fever (pyrexia) which is a consistent clinical finding in all CSF infections (Moennig et al., 2003). These initial symptoms are generally followed by vomiting, diarrhea, labored breathing (dyspnoea) and coughing. Invariably animals that contract the acute form of the disease develop central nervous system disorders which are evident by the development of a staggering gait, weakness of the hind quarter and incoordination (ataxia) which eventually progresses to the point where the animal is unable to stand at all. In the terminal stage of disease, the infected animal typically becomes completely unresponsive and may lie on the ground continually moving its limbs, an action commonly referred to as peddling. Death usually occurs within 5 to 9 days after infection (van Oirschot, J.

T., 1986).

The chronic form of CSF infection differs from the acute form in that neurological symptoms are not observed and there is a longer duration of infection with death occurring between 3-4 weeks. Over this course of time, animals begin wasting, losing both body mass and strength and commonly develop cyanosis of the skin in its extremities, beginning in the ears and proceeding to the snout, tail and eventually to the legs and ventral region. (van Oirschot, J. T.,

1986). In addition, chronically infected animals normally become severely immunosuppressed and develop secondary infections such as bronchopneumonia. Occasionally animals that contract the chronic form of disease recover from the disease developing antibodies which protect it from future CSF infection.

The last form of CSF infection is referred to as late onset and is a congenital disease in piglets. CSFV has a tropism for the reproductive tissues and is readily able to cross the placenta in pregnant sows. Infection during early pregnancy typically results in abortion however infection in later stages (50-70 days) of pregnancy can lead to the birth of persistently viremic piglets. At birth the piglets appear clinically normal but as they mature they exhibit poor growth

(runting), wasting, and congenital tremors. Passive immunity provided by maternal antibodies generally protects piglets against mortality during the first 5 weeks of life, but not against the replication and shedding of virus (van Oirschot, J. T., 1986). The survival period of infected piglets is usually a few months.

Upon post mortem examination, animals infected with CSFV generally present a widespread petechiae of the skin, mucous membranes, lymph nodes, laranyx, bladder, kidney, and ileum (van Oirschot, J. T., 1986). The most noticeably affected organs are the lymph nodes,

spleen, and kidneys. The lymph nodes commonly appear enlarged, edematous and hemorrhagic while the spleen may exhibit multifocal infarctions which appear as variably sized dark blebs found on periphery and apex of the spleen. Lesions of the kidney also range in size from petechiae to ecchymotic hemorrhages. In addition, hemorrhagic lesions may be found on the liver, lungs and heart. A generalized depletion of lymphoid tissues is often observed in final stages of disease along with the presence of a non-purulent encephalitis particularly in animals that had contracted the acute form of the disease (van Oirschot, J. T., 1986).

Importantly, analysis of blood samples drawn from infected pigs reveals yet another prominent manifestation of CSF infection which is the development of a severe leukopenia (low white blood cell count) and marked thrombocytopenia (decreased platelet count). The development of these conditions are invariably the causal factor for characteristic symptoms observed, with the leukopenia probably being the underlying cause for the immunosuppression seen during chronic courses of infection while the thrombocytopenia can account for the generalized hemorrhagic nature of the disease. Another interesting finding upon analysis of clinical samples taken from infected animals is the extent of viral dissemination throughout the entire body with virus being able to be isolated from all of the major organs as well as blood.

Pathogenesis

Under natural conditions, CSFV enters the host through oral, nasal, conjunctival and genital mucous membranes (Leiss, 1987). The virus initially infects epithelial cells of the

tonsillar crypts (Van Oirschot et al., 1989). The virus then travels via the lymphatic vessels to the lymph nodes draining the tonsils. The virus subsequently replicates in the regional lymph nodes and eventually enters the blood producing a primary viremia. Monocytes are early target cells for the virus. Accordingly, the virus carried by these cells through the peripheral bloodstream allowing the virus access to its primary targets readily infecting the spleen, bone marrow, visceral lymph nodes, and lymphoid structures lining the small intestines (Paton et al., 2003).

The virus multiplies in these tissues of the reticuloendothelial system reaching very high titers, upon which the virus is released into bloodstream again associating monocytes producing a secondary viremia (van Oirschot, J. T., 1986). As a result of the monocytes ability to migrate into virtually every organ of the body, the secondary viremia disseminates the virus to the remaining parenchymateous organs.

The time in which virus spread is completed mainly depends on the virulence of the virus. Highly virulent virus can be detected in most organs by 5-6 days PI. Importantly, CSFV has an affinity for reticular endothelial and epithelial cells. Resultantly, widespread hemorrhages are observed in acute CSF due to degeneration and necrosis of endothelial cells, as well as thrombocytopenia and disturbances in fibrinogen synthesis (Pauly et al., 1998; Gómez-

Villamandos et al., 2001). Consequently, acute CSFV is almost always fatal.

Viral Transmission and Route of CSFV Infection

Domestic and wild pigs are the only natural reservoir for CSFV. The primary route of viral transmission is by direct contact between infected and susceptible pigs. Infected pigs excrete virus in oral, nasal, and lacrimal secretions as well as urine and feces (van Oirschot, J. T.,

1986). The virus gains entry into a susceptible animal by ingestion and/or inhalation of any of these bodily fluids, as well as contact with the conjuctiva, genital mucous membranes or skin abrasions. The virus may also be spread by indirect means via contaminated transport vehicle or by persons such as veterinarians, farmers, and other animal personnel and their equipment. In addition, it has been suggested that rodents, pets, and birds could also serve as possible mechanical vectors. Transmission of CSFV via these vectors is believed to occur by the vector becoming contaminated through contact with excretions of infected pigs and thereby providing transportation for the virus particles to other susceptible pigs. Further sources of infection include the feeding of improperly sterilized swill, artificial insemination with semen from infected swine, and contact with wild boar that may be infected. This is especially true in Europe where CSFV is known to be endemic in the wild boar population (Edwards et al., 2000).

Epidemiology

The Office of International des Epizooties (OIE: world organization for animal health) classifies CSFV as a notifiable disease. All notifiable pathogens are defined by the OIE as

transmissible diseases that have the potential for very serious and rapid spread, irrespective of national borders, that are of serious socio-economic and/or public health consequence and are of major importance in the international trade of animals and animal products (OIE, 2016).

Moreover, the OIE considers CSF the most economically damaging swine disease worldwide.

The first recognized outbreak of CSF was documented in Ohio in 1833, however it remains speculative as to whether this U.S. city was indeed the initial source of the disease or whether Europe was the actual origination of the disease. This latter belief is supported by reports of an epizootic resembling CSF in France as early as 1822 as well as the reappearance of the epizootic in Germany in 1833 (van Oirschot, J. T., 1986). Regardless of the origin, it is known that by the early 1860s CSFV was widespread throughout both Europe and America

(Edwards et al., 2000).

At the present time CSF is endemic in much of Asia, Eastern Europe, Central and South

America, the southern states of Mexico (Fig. 3). Outbreaks have also been reported in the

Caribbean nations of Cuba, Haiti and the Dominican Republic. The situation in Africa is largely unknown however CSF has been reported in Africa’s island nation of Madagascar. Regions currently believed to be free of CSF include Australia, New Zealand, United States, Canada, and

Western Europe.

CSF was eradicated from the United States in 1978 after a 16-year effort by the farm industry and State and Federal governments. Of particular concern to the United States as a CSF free nation is the presence of CSFV in both Mexico and the Caribbean. The close proximity of these countries to the U.S. makes them a potential reservoir of infection and imminent threat for the reintroduction of CSFV into the United States.

Pathways for CSFV entry into US

Intentional Release of CSFV

There are many pathways in which CSFV could be reintroduced into the United States which may occur either by intentional or unintentional release. Intentional release may occur by the weaponization of viral diseases by foreign terrorist cells targeting livestock. There is substantial evidence that terrorists have considered culturing biological weapons for this purpose.

For example in early 2002, United States Navy SEALs stormed a cluster of caves in eastern

Afghanistan, known to be an Al Qaeda storehouse expecting to find guns and explosives.

Instead, they found hundreds of documents all planning a terrorist attack on the United States food supply. There were agriculture articles from American science journals which were translated into Arabic. A comprehensive list of the most devastating livestock pathogens such as

FMDV, CSF and rinderpest was also found. Most alarmingly, was the discovery of training documents, detailing how to deploy these pathogens on farms. Furthermore, in 2002 CIA agents found a dossier containing information on Plum Island Animal Disease Center which studies

foreign animal diseases such as FMD, CSFV, and ASFV, in a raid of Sultan Bashiruddin

Mahmood’s residence in Afghanistan. Mahmood was a nuclear physicist from Pakistan and a known associate of Osama bin Laden. Lastly, in 2008 Aafia Siddiqui, aka Lady Al Queda, a

Pakistan neuroscientist and MIT graduate, as well as a relative of 9/11 mastermind Khalid

Sheikh Muhammad was arrested in Afghanistan. At the time of her apprehension among the items found in her possession were hand-written notes about a "mass-casualty attack" and a list of targets which included Wall Street, Brooklyn Bridge, Statue of Liberty, Empire State Building and the Plum Island Animal Disease Center.

Unintentional Release of CSFV

The risk of introduction of CSFV through unintentional acts would mostly likely involve indirect exposure of a host through a formite, with a formite constituting nearly any object that could potentially become contaminated with CSFV such as contaminated meat, boots, or clothing just to name a few. There are several different avenues in which a contaminated formite might enter the U.S. For instance, today’s ease of international travel has introduced a new means for diseases to be disseminated globally. The current volume and speed of travel has increased in industrialized nations at a rate of more than 1000-fold in the last century (Wilson,

2003). Consequently, reintroduction of CSFV could result from travelers entering the U.S. from endemic countries with contaminated items. To serve as an illustration of the volume of travelers entering the US, in 2015 there were over 75 million international travelers that visited the country. Many of these visitors came from regions endemically infected with CSFV, 9.7 million were from Asia, 1.2 million from the Middle East, 5.5 million from South America, 1.3million 50

from the Caribbean and 17 million from Mexico (http://travel.trade.gov/view/m-2014-I-

001/index.html). Of particular concern of these figures is the volume of travelers coming from

Mexico where CSFV is believed to be endemic in the pig population of backyard farmers.

Exceedingly, the immense number of annual international visitors is compounded by the number of immigrants present in the country. In 2014, there were 42.4 million total immigrants with 11.7 million originating from Mexico (migrationpolicy.org). Furthermore in 2014, it was estimated that there was 11.4 million illegal nationals residing in the US. The significance of these statistics involving foreign born residents is that it is presumable that some portion of this population will receive mail and packages from their family and friends remaining in their native countries. In addition, cultural preferences would likely result in this population receiving food products from countries where they were born. Resultantly, there is a possibility that some of these items may have potentially been exposed to CSF virus which serves as another way for virus to enter the US. If any of these items coming into US were infected with CSFV then an outbreak could occur assuming that the item came into contact with a susceptible animal.

Lastly, CSFV exposure could also occur through direct contact of contagious animals arriving from outside the US with domestic livestock. Notably, the United States imported 5.8 million live hog in 2011. The majority of these pigs, specifically 79 percent, were from Canada.

While, Denmark supplied 10% of these pigs to the U.S. (http://www.agmrc.org/commodities- products/livestock/pork/pork-international-markets-profile). Although the risk of animals entering the US from CSFV free countries and infecting domestic livestock is minimal, the US

agricultural and customs officials must remain vigilant as the status of these countries could change at any time.

CSFV Control Strategies

The OIE is responsible for setting internationally agreed standards relating to animal health and conditions under which a country can be considered free of CSF. Due to the highly infectious nature of CSF, disease free countries have introduced trade barriers that prohibit importation of animals or animal products from countries that do not possess CSF free status with the ban continuing until the OIE recertifies that country as CSF free.

Currently, there are two strategies utilized in the prevention and control of CSF. Usage of either strategy results in international trade restrictions of pork and pork products for a set period of time as outlined by the OIE. The first strategy is the use of traditional LAVs which are commonly employed in regions where CSF is endemic, such as, Asia and South and Central

America. Live attenuated CSFV vaccines have been in existence since the 1960s with the most widely used CSFV LAV being the Chinese (C) Strain. CSFV LAVs are avirulent strains of

CSFV, which have been attenuated by adaptive mutations obtained through serial passage in either rabbits, as is the case for C strain, or passage in tissue cell culture. In general, modified live attenuated vaccines are easy and economical to produce. These vaccines replicate in the host without causing significant adverse clinical effects, and provide protection against challenge with

virulent virus of the same serotype typically after the administration of just one infectious dose.

The major advantage that this type of vaccine has over others is that it is able to elicit both cellular and humoral immune responses, which are able to protect host animal against infection with wild type virus. Accordingly, CSFV LAVs have been proven both safe and efficient providing protection in animals as early as 7 days after vaccination (Terpstra et al., 1990).

However, the drawback of using live attenuated CSFV vaccines is that vaccinated animals cannot be serologically differentiated from infected animals. As a result, once emergency vaccination is implemented in a region, that region is thereafter prohibited from international trade for a period of at least 2 years before the OIE will recertify that area with a CSF free status

(van Oirschot et al., 2003). The resulting ban on pork and pork products could potentially result in huge economic losses for the affected region.

The second strategy for CSF control is a non-immunization “stamping out” policy which has a more expedient time frame in the return to CSF free status. Due to the economic benefits of rapid return to disease-free status, the European Union (EU) and some parts of Eastern Europe have opted for this latter approach. The “stamping out” policy has been in effect in the EU since

1980 and was instituted in an attempt to eradicate CSF as well as to ensure free trade of pigs within the EU member states.

Under the stringent regulations of the “stamping out” policy, all animals on a farm where

CSF is detected must be destroyed. In addition, all pigs on farms within a 3 km radius of a CSF infected farm must be pre-emptively slaughtered, with this area being referred to as the protection zone (Stegeman et al., 2000). Finally, farms 7 km beyond the protection zone would

be established as a surveillance zone and a movement ban would be enforced for these farms for a period of at least six months while the farms in this area are periodically tested for the presence of CSF (Stegeman et al., 2000). If no additional CSF cases are detected over this time the OIE will restore a CSF free status to the region.

The large scale culling in the surveillance zone is very problematic in that many healthy animals eventually have to be destroyed over the 6 month surveillance period due to welfare reasons caused by the overcrowding of farms and the lack of income to feed animals on overcrowded farms (Stegeman et al., 2000). An outbreak of CSFV in the Netherlands exemplifies the serious economic ramifications that this large scale culling can cause. The

Netherlands outbreak was the largest and most devastating of any outbreak that has occurred in

Europe since the implementation of the “stamping out” policy. The outbreak began in February of 1997 and progressed into an uncontrollable epidemic that lasted until March of 1998. During this period 429 CSF-infected herds (700,000 pigs) were destroyed and 1286 herds (1,125,000 pigs) were pre-emptively slaughtered (Stegeman et al., 2000). Incredibly, an additional

10,567,000 pigs not infected with CSF had to be killed for welfare reasons. (Stegeman et al.,

2000). The total economic loss that the Netherlands endured during this epidemic amounted to

2.3 billion in U.S. currency.

Due to the huge economical losses caused by the non-vaccination policy as well as ethical objections to the mass killing of healthy animals, modifications to the current policy are needed. One option that has been given high priority is the development of a safe “marker” or

differentiating infected from vaccinated animal (DIVA) vaccine with an accompanying discriminatory antibody assay that would allow for the differentiation of infected animals from vaccinated animals (van Oirschot et al., 2003). The use of such a DIVA vaccine would be used for emergency vaccination in conjunction with the “stamping out” strategy during a CSFV outbreak to limit the duration and extent of the outbreak, as well as, minimize the economic losses caused by the outbreak by reducing the number of pigs that have to be destroyed. In addition, an efficient marker vaccine would also be useful in the implementation of control programs in countries where the disease is endemic.

Live Viral Vector Vaccine Candidates

Over the last two decades various strategies have been utilized to develope experimental vaccines in an attempt to satisfy the need for safe and efficient DIVA vaccines. Early efforts were focused on the use of recombinant live attenuated viral vectors such as vaccinia virus, pseudorabies virus, and porcine adenovirus. These live attenuated viruses, which are harmless to swine, are able to act as delivery vectors expressing its inserted foreign immunogenic antigen(s).

The benefit of using live viral vectors as vaccines is that live vaccines are excellent inducers of the immune system because they are able to replicate in cells eliciting both humoral and cellular responses against the inserted antigen thereby mediating protection in pigs against CSFV. Since only a portion of the CSFV genome would be included in the viral vector the resulting antibody pattern in an immunized animal would substantially differ from that of a CSFV infected animal.

The first two CSFV viral vector vaccines to be developed were recombinant vaccinia viruses, with one vector expressing all four CSFV structural proteins; C, E0, E1 and E2 and the other vector similarly expressing the structural proteins but lacking most of the E2 glycoprotein

(Rumenapf et al., 1991). Notably, both vaccine candidates were found to mediate protection in pigs against lethal CSFV challenge at 42 days post immunization. Subsequently, the next recombinant viral vector to be utilized as a CSFV vaccine was the Pseudorabies virus vector containing only the E2 glycoprotein. Immunization with this experimental vaccine was also able to confer protection in swine at 42 days (van Zijl et al., 1991). Similar results were later reported for expression of E2 in the vaccinia virus vector (Konig et al., 1995) as well as in the porcine adenovirus vector (Hammond et al., 2000), although, in both of these cases protective immunity was established a full week earlier. In addition, vaccination with vaccinia virus expressing the E0 glycoprotein of CSFV was also found to protect swine against virulent challenge at 35 days

(Konig et al., 1995).

Although animals immunized with these live recombinant virus vectors successfully acquired protective immunity against CSFV, a major deterrent in using these recombinant vaccine candidates is the pathogenic potential of the vector viruses to non-vaccinated humans as well as animals. As a result, the use of live viral vector recombinant vaccines is currently prohibited.

Non-replicating Subunit Vaccine Candidate

The unsuitability of viral vectors for development of recombinant vaccine candidates spurred the necessity for further research into safer vaccine alternatives. To this end, one successful option considered was a protein based non-replicating subunit vaccine. Unlike inactivated vaccines, non-replicating subunit vaccines contain only the immunogenic antigen portion of a pathogen, which is necessary to elicit a protective immune response. Identifying which antigens best stimulates the immune system is a complicated and time-consuming process.

Non-replicating subunit vaccines have a general safety advantage over live attenuated vaccines due to the fact that they are non-living, non-replicating molecules, as such, there is absolutely no risk of inducing disease. Furthermore, subunit vaccines have an excellent stability profile and are more stable than live attenuated viruses. However, in contrast to live vaccines where the immune response closely resembles natural infection, the immune response to a non- replicating subunit vaccine is mostly humoral with little to no cellular immunity raised. Due to this, the first dose of a non–replicating subunit vaccine does not usually produce protective immunity, but merely

“primes” the immune system. It is the second or third dose, referred to as a booster shot which stimulates a protective immune response. Another disadvantage of the non-replicating subunit vaccine is that antibody titers against inactivated antigen(s) diminish with time. As a result, non- replicating subunit vaccines may require intermittent immunization with booster shots to increase antibody titers to maintain active immunity to the desired antigen(s).

The most immunogenic of both the structural and non-structural CSFV proteins is the E2 envelope glycoprotein. Expression of the E2 protein alone has been established to be sufficient to

raise high levels of neutralizing antibodies and confer protective immunity in swine against

CSFV. For these reasons, the E2 glycoprotein protein has been the target of CSFV non- replicating subunit vaccine research. Notably, the successful non–replicating subunit vaccine candidate that has been developed contains E2 produced in insect cells by a baculovirus expression vector (Van Rijn et al., 1996). The advantage of using this expression system is that insect cells are able to glycosylate proteins, as such the resulting viral E2 glycoprotein is glycosylated and in so doing is thought to be expressed in a more ‘natural’ way. The E2 proteins produced in the baculovirus system are subsequently adjuvanted with a water–oil–water emulsion for final non-replicating subunit vaccine preparation (Moormann et al., 2000).

When the E2 non-replicating subunit vaccine was tested in vaccination/challenge experiments in swine, it was found that a single dose of the non-replicating subunit vaccine was unable to produce protection at 7 days post lethal challenge nor 10 days post challenge (Bouma et al., 2000 and Uttenthal et al., 2001). In fact, complete protection was not observed until 14 days post challenge with protection lasting up to 13 months (Bouma et al., 2000). However, marked improvement in immunity was observed when a second dose of non-replicating E2 subunit was administered with clinical protection developing by 7 days post challenge (Dewulf et al., 2001).

The E2 non-replicating subunit vaccine was the first to be considered a marker vaccine as discrimination between vaccinated and infected pigs can be based on the detection of antibodies against Erns and/or NS3 proteins (Moormann et al., 2000). Importantly, after infection with

CSFV, antibodies are raised against two structural glycoproteins, E2 and Erns, and one non-

structural protein NS3 (Terpstra et al., 1988; Paton et al., 1991; Kwang et al., 1992). As would be expected, vaccination with non-replicating subunit E2 vaccine only raises antibodies against

E2. Therefore, the detection of specific antibodies by an ELISA assay would be able to provide a direct measure as to the immune status of animals and establish whether animals have been vaccinated or CSFV infected. Resultantly, the accompanying ELISA test for the E2 non- replicating subunit was based on the detection of Erns glycoprotein antibodies. Detection of NS3 protein antibodies was not utilized due to the fact that the NS3 protein is conserved among pestiviruses (Moorman et al., 2000). Unfortunately, there are limitations with the anti-Erns

ELISA also. First, antibodies against Erns do not develop until 3–6 weeks after CSFV infection which makes early detection unfeasible (de Smit et al., 2000). Second, only one of 3 different anti-Erns ELISAs developed was found to be specific for CSFV, the others two were found to cross-reactive with Erns antibodies from other pestiviruses. Consequently, it was concluded that the currently available anti-Erns ELISA assays are not sensitive or specific enough to detect

CSFV infected animals reliably within vaccinated herds. Both the lack of CSF-specificity of

Erns-ELISA assays combined with the need for several injections makes the E2 non-replicating subunit marker vaccine unsuitable for use as an emergency vaccination to control a CSFV outbreak.

Recombinant Pestivirus Vaccine Candidates

The advent of reverse genetics technology has enabled researchers to construct and genetically modify cDNA infectious clones (IC) of CSFV strains in order to produce live

attenuated pestivirus vaccines. Live attenuated vaccines are usually the most efficient of all the different types of vaccines in raising a protective immune response as they have the most similar mode of infection as the wild type virus. Several novel reverse genetic approaches have been employed over the last two decades to manipulate cDNA IC in an effort to identify virulence factors and produce live attenuated CSF viruses. Resultantly, a wide array of virulence factors have been identified in the CSFV genome. Importantly, various genetic mutations in the proceeding CSFV genes in the context of virulent CSFV IC have all been demonstrated to be effective mechanisms of attenuation: Npro, Core, Erns, E1, E2, p7, and NS4B. Moreover, virus mutants containing most of these mutations have also been shown to provide protection in pigs against lethal challenge.

One approach that was utilized to generate a novel live attenuated CSF virus was by deleting the Npro gene from the CSFV genome. It had previously been demonstrated that the

Npro gene was dispensable for CSF viral replication by replacing the coding sequence for Npro in the full length infectious clone of CSFV strain of Alfort/187 with nucleotide sequence for the murine ubiquitin gene (Tratschin et al., 1998). Subsequently, this group later tested their Npro deletion viruses in pigs for attenuation and potential to provide protection from CSF (Mayer, et al. 2004). They observed that removal of Npro in the moderately virulent Alfort and highly virulent Eystrup CSFV strains was able to attenuate the virulent phenotype of both parental strains as well as protect against challenge with a lethal dose of highly virulent CSFV at 21 days post immunization.

Another method researchers have employed to produce live attenuated vaccines is by producing chimeric viruses via substituting gene(s) of one virus with the corresponding gene(s) from a closely related virus or strain. Accordingly, two full length chimeric viruses were constructed in which either the E2 or the E0 antigenic regions of BVDV II strain 5250 were substituted for the homologous regions in full length infectious clones of CSFV C strain vaccine

(van Gennip et al., 2001). Importantly, the use of the BVDV antigenic regions in these constructs would allow for the potential serological discrimination between vaccinated and naturally infected animals. When evaluated in animals both of the resulting chimeric viruses continued to exhibit the attenuated phenotype of the parental C strain as well as conferred protection against lethal CSFV challenge at 28 days post infection. Unfortunately, neither of these CSFV/BVDV chimeric viruses were able to fully prevent transmission of the challenge CSFV virus from vaccinated animals to sentinels (de Smit et al., 2001). Consequently, another potential chimeric virus with marker capabilities was constructed, this time however using an infectious cDNA clone of a BVDV vaccine strain, CP7, in which the E2 gene in the BVDV IC was replaced with the analogous region from CSFV strain Alfort/187 (Reimann et al., 2004). As expected this chimeric virus was found to be attenuated in animals as the BVDV backbone harboring CSFV

E2 is apathogenic in pigs. Importantly, this chimeric virus provided complete protection in swine when challenged with virulent CSFV at 28 days after infection. The implications of the attenuation and protective immunity generated by this chimeric virus coupled with its DIVA capability will be discussed further in the next section.

While these putative marker vaccine candidates were being developed, another research group was using the same chimera technology to detect CSFV virulence factors and host range determinants in an effort to use the knowledge gained in these areas to rationally design putative

attenuated marker vaccines. To accomplish this objective, a series of chimeric viruses were initially constructed where regions of CSFV genome were systematically replaced with the homologous regions of attenuated vaccine CSFV strain CS in the genetic backbone of highly virulent CSFV Brescia strain (Risatti, et al., 2005a). The resulting chimeric viruses were evaluated in swine for viral virulence. Interestingly, only chimeras harboring the CS E2 gene were attenuated suggesting that a significant virulence determinant absent in CS E2 was responsible for attenuation. As such, a separate chimeric virus was produced in which only the

E2 genomic region of the pathogenic Brescia strain was replaced with analogous region from the

CS vaccine strain (Risatti et al., 2005a). When this chimeric virus was tested in swine, a significant in vivo attenuation was observed. Importantly, animals immunized with this virus were also found to be protected from lethal challenge as early as 3 days post vaccination. Further research identified the exact amino acid residues in CS E2 producing the attenuation of Brescia virus (Risatti et al. 2006). In addition, it was found that attenuation of Brescia coincided with the elimination of reactivity associated with a specific epitope residing in CSFV E2 that is recognized by monoclonal antibody (mAb) WH303. Notably, mAb WH303 is routinely used for

CSF diagnostics due to its inability to react with the E2 region from related pestiviruses, BVDV and BDV (Lin et al., 2000). The amino acids responsible for binding mAb WH303 were determined by progressive mutations introduced into an area of E2 between residues 829 and

837. The corresponding amino acid sequence, TAVSPTTLR, of CSFV Brescia was progressively replaced with the corresponding amino acid residues of BVDV, TSFNMDTLR, producing 4 chimeric viruses. The resulting viruses were designated T1v – T4v and were evaluated in animals. T1v – T3v induced lethal disease, while T4v, containing BVDV residues

TSFNMD, induced mild transient disease in pigs and culminated with the elimination of

reactivity with mAb WH303. Remarkably, animals infected with T4v were protected when challenged with virulent Brescia virus at 3 and 21 days post-vaccination implicating the homologous CSFV residues, TAVSPT, as a significant determinant of CSFV virulence.

In addition, researchers have successfully generated several live attenuated viruses by targeting known protein functions associated with specific CSFV genes by mutating various amino acids in the virulent genomic sequences of different CSFV strains. As a result, a number of virulence factors in CSFV have been identified. For instance, in a study designed to abrogate the RNase activity of Erns in an effort to determine the function of this enzyme on CSFV pathogenesis and virulence, researchers constructed multiple mutants substituting either lysine or leucine at two specific residues, H297 and H346, in the E0 gene of CSFV strain Alfort/187

(Meyers et al. 1999). Notably, these targeted residues were theorized to be involved in catalytic reaction of RNase activity (Meyers et al., 1999). Animal experiments with the resulting virus variants showed that all of the mutations inactivating the RNase had an attenuating effect in swine. Although the degree of attenuation varied among the variant viruses, all of the infected animals effectively developed protective immunity, as demonstrated by the challenge with highly pathogenic CSFV Eystrup 10 weeks after the immunization implicating RNase activity in Erns as one of many determinants of virulence in CSFV that will be discussed throughout this chapter.

Similarly, glycosylation sites in all three CSFV glycoproteins have been found to influence viral virulence as mutations of several individual N-linked glycosylated sites residing in Erns, E1 and E2 of highly virulent Brescia resulted in attenuation of the parental virus in pigs

(Fernandez-Sainz, et al. 2008 ; Fernandez-Sainz et al., 2009; Risatti et al., 2007). To briefly sum up the findings from these studies, mutation of one glycosylation site, N269A, in Erns, substitutions in three glycosylation sites in E1, N594A and N500A/N314A and alanine exchanges in two N-linked glycosylation sites residing in E2, N116A and N185A, produced attenuated phenotypes in vivo. Notably, each of the corresponding nonglycosylated mutant viruses were also able to elicit protection against lethal CSFV challenge as earlier as 3 days post- inoculation.

CSFV intermolecular disulfide bonds have also been determined to be factors of virulence in the homodimerization and heterodimerization of glycoproteins Erns and E1, respectively (Tews et al., 2009; Fernandez-Sainz et al., 2011). Importantly, the formation of disulfide bridges is mediated between cysteine residues. It has been shown that eight of the nine conserved cysteines in glycoprotein Erns form intramolecular disulfide bonds while a ninth cysteine residue found near the carboxyl end at residue 171 of the Erns protein is responsible for formation of Erns homodimers (Langedijk et al., 2002). Deletion or replacement of amino acid

C171 to either Phe or Ser results in loss of dimerization as revealed by immunoprecipitation and western blot (Tews et al. 2009). Furthermore, a mutant virus containing substitution, C171S, in infectious cDNA clone of CSFV Alfort/Tübingen was markedly attenuated in pigs. Likewise, the

E1 envelope protein contains six cysteine residues. The role of these E1 cysteine residues in formation of E1-E2 heterodimers and resulting effect on CSFV replication and virulence in swine was determined by generating a panel of single and double recombinant mutant viruses substituting cysteine residues with serine in E1 glycoprotein (Fernandez-Sainz et al., 2011). Only one recombinant virus in the panel of mutants resulted in an attenuated phenotype in pigs with

that virus containing two amino acid substitutions, C24S and C94S, in E1 glycoprotein suggesting that these residues are critical in the proper formation of E1-E2 heterodimers and virulence in swine. In addition, the attenuated mutant displayed a partial disruption of E1-E2 heterodimer formation in infected cells as well as decreased growth rate in primary swine macrophages. Despite its diminished growth in vitro, this mutant virus provided protection in swine challenged with virulent CSFV at 3 and 28 days post infection. These findings strongly suggest that Erns and E1-E2 dimerization are virulence determinants of CSFV and prevention of

Erns and E1-E2 dimerization leads to attenuation of CSFV.

Structural prediction analysis software has also been effectively utilized to identify specific domains in p7 and NS4B involved in viral virulence. Previously, the p7 protein of hepatitis C, HCV, has been characterized as a member of viroporin family and found to be crucial for the production and release of infectious HCV particles from infected cells (Griffin et al., 2003). Viroporins by definition are small, highly hydrophobic, virus-encoded proteins that able to oligomerize to form hydrophilic ion channels in host cell membranes (Scott, et al., 2015).

Due to structural similarities between the p7 protein of HCV and CSFV it has been suggested that CSFV p7 may similarly function as a viroporin involved in pore formation and release of infectious particles across cell membranes. To test this premise, a series of p7 recombinant mutant viruses each containing either three to six consecutive alanine substitutions spanning the entire protein was created using a scanning mutagenesis approach (Gladue et al., 2012). These recombinant viruses resulted in the identification of regions within CSFV p7 that are critical for virus production in vitro and resultantly demonstrated that the C-terminal transmembrane helix of p7 does in deed possess pore-forming activity. While, some of the virus constructs where

unable to produce infectious progeny, other constructs produced viruses that were either partially or completely attenuated in animals, indicating that CSFV p7 plays a significant role in viral virulence.

Similarly, a Toll/interleukin-1 receptor (TIR)-like domain has been determined to reside in NS4B protein. This domain is unique to both CSFV and BVDV and is not found in other pestiviruses or flaviviruses. TIR domains are known to participate in signal transduction of Toll- like receptors (TLRs). In general, TLRs are a large family of proteins that play a crucial role in recognition of various microbial components, activate the innate immune system, and induce of cytokines needed for the adaptive immune response (Akira et al., 2004). Importantly, the TIR domain contains 3 conserved motifs, termed box 1, 2 and 3. It has been shown that alanine substitutions in box 1 and 2 of a TIR domain in a human interleukin receptor can inhibit its signaling, while signaling was unaffected by alanine substitutions in box3 of the TIR domain.

Based on these observations, Fernandez-Sainz et al. (2010) selectively generated mutations in boxes 1 and 2 in order to elucidate the function of this TIR domain in CSFV NS4B. Mutations,

V2566A, G2567A, and I2568A in the box 2 domain produced a virus (NS4B VGIv) with an attenuated phenotype in pigs. Infection with recombinant NS4B VGI was characterized by localized replication in the tonsils with limited viral dissemination during infection. Furthermore, animals inoculated with NS4B VGIv were completely protected against lethal CSFV challenge as early as 3 days post infection. Analysis of cellular gene expression of infected primary swine macrophage revealed cultures infected with NS4B VGIv showed increased activation of several genes, compared to those in cultures infected with the parental virus. Notably, most of the genes exhibiting increased transcriptional activation with NS4B VGIv infection are similarly induced

by activated TLR-7 (Lee et al., 2003). Furthermore, TLR-7 functional involvement was indirectly confirmed by the observation that transfection of macrophages with wild-type NS4B reduces the responsiveness of TLR-7 to a well characterized activator, imiquimod R837 (I-R837) while transfection with NS4B gene harboring box 2 domain mutations was less efficient at mediating this reduction. Fernandez-Sainz et al. (2010) speculated that these findings suggest that NS4B of wild-type CSFV blocks TLR-7 transcriptional activation during infection in animals.

In addition, using proteomic computational analysis, a putative fusion protein (FP) was suggested to reside between amino acid residues 818CPIGWTGVIEC828 of CSFV E2 (Garry and

Dash, 2003). To discover the role of this putative FP on in vitro and in vivo replication of CSFV as well as in the production of disease in swine, a series of recombinant CSF viruses containing amino acid substitutions in the FP area were designed using the cDNA infectious clone of the virulent CSFV Brescia strain (BICv) (Fernandez-Sainz et al., 2014). Resultantly, none of the mutations resulted in an attenuated phenotype in animals but double (C818S/C828S) and triple

(P819S/I820S/W822S) residue substitutions were found to completely abrogate virus replication.

Subsequently, synthetic peptides representing native, double and triple mutant FP variants were generated and assessed for their ability to penetrate into lipid bilayers. While no significant alterations to the secondary structure were observed among the three synthetic FP, the double and triple mutations did appear to affect the degree in which the peptide was able to insert itself into model membranes. These findings reveal that amino acid sequence, 818CPIGWTGVIEC828, is critical for membrane fusion and virus replication.

More recently, yeast two hybrid system has been employed to identify viral/host protein- protein interactions for CSFV as another tool to investigate viral virulence. Yeast two hybrid screening revealed several swine proteins which specifically bind to CSFV Core protein; Sumo-

1, UCB-9, IQGAP-1 and OS9. Of these host proteins, SUMO-1 and UBC9 are both involved in the cellular SUMOylation pathway (Niedenthal, 2007). Importantly, several viruses are able to effectively evade the host immune response by inducing modifications to the host SUMOylation pathway (Sadanari et al., 2005; Huh et al., 2008; Shirai et al., 2008; Chiu et al., 2007; Yueh et al., 2006; Palacios et al., 2005). As such specific residues within CSFV Core protein mediating the interaction with SUMO-1 and UBC9 proteins were identified (Gladue et al., 2010). Amino acid substitutions of these residues to alanines in the viral genome resulted in recombinant viruses with attenuated phenotypes indicating that CSFV virulence is associated with the ability of Core protein to interact with host proteins in the cellular SUMOylation pathway.

Subsequently, interaction of the cellular protein, IQGAP1 with CSFV core was examined to determine its significance (Gladue et al., 2011). IQGAP1 protein has been shown to play an important role as a regulator of the actin cytoskeleton (Fukata et al., 2002; Watanabe et al.,

2004). Notably, it has been reported that interaction of IQGAP1 with the Moloney murine leukemia virus (MMLV) matrix (M) protein plays an important role in the intracellular trafficking of MMLV, which is essential for virus replication (Leung et al., 2006). In this same study the critical residues within MMLV M protein that interact with IQGAP1 were mapped by mutagenesis experiments. Based upon these findings, four regions within CSFV Core, designated

I – IV, were identified that potentially bind IQGAP1. (Gladue et al., 2011). Amino acid residues in these putative binding sites were replaced with alanines. Resulting mutations in areas I and III completely disrupted the Core–IQGAP1 protein–protein interaction while substitutions in areas

II and IV only partially affected protein binding. CSFV mutants harboring substitutions in these four areas were assessed in animals to determine importance of the Core–IQGAP1 protein interaction on virulence. Interestingly, substitutions in regions I and III of the Core, resulted in complete attenuation in swine. These results suggest that the ability of CSFV Core protein to bind cellular IQGAP1 protein during infection plays a critical role in virus virulence within the swine host. Lastly, osteosarcoma amplified 9 protein (OS9) a protein involved in the endoplasmic reticulum-associated degradation (ERAD) pathway, was identified by YTH system to interact with CSFV Core (Gladue et al., 2014a). OS9 has been shown to be involved in the degradation of misfolded glycoproteins (Huttner et al., 2012), which could be significant considering CSFV Erns, E1 and E2 are heavily glycosylated structural proteins. Therefore, it is possible that binding of OS9 by the Core protein could prevent degradation of viral glycoproteins. Utilizing alanine scanning mutagenesis, the OS9 binding site within the CSFV

Core protein was mapped to the C-terminus of unprocessed Core, before its cleavage by signal peptide peptidase. A recombinant CSFV mutant lacking OS9 binding site in Core protein presented a significant decrease in its ability to replicate in primary swine macrophage cell cultures, the natural cell target of CSFV. However, no differences were observed in mutant infected swine and animals infected with parental CSFV. As a result, it was concluded that the

OS9-Core interaction affects the efficiency of virus replication by an unclear mechanism.

Overall, the YTH system and resulting identification of Core host proteins partners has provided interesting insight into the pathogenesis of CSFV by elucidating what appears to be multiple roles for Core in the CSFV replication cycle.

Lastly, a transposon linker insertion mutagenesis approach was also utilized to identify virulence and host range determinants of CSFV. Hence, Risatti et al. (2005b) used transposon linker insertion mutagenesis (TLIM) to mutate an infectious clone of the pathogenic strain

Brescia, BIC (Risatti et al., 2005b). Significantly, one of the resulting viral mutants with an in- frame transposon linker insertion of 57 nucleotides in the carboxyl terminal domain of E1, at genomic position 2429, four residues upstream of the cleavage site between E1 and E2, was found to attenuate the Brescia virus thus identifying a virulence determinant associated with the

E1 glycoprotein of CSFV. Infected animals exhibited decreased replication in tonsils, limited generalization of infection, and significant reduction of viral shedding. Notably, these animals survived lethal challenge of CSFV presenting no signs of clinical disease. These results indicate that this mutation in E1 glycoprotein attenuates the virus and consequently has been identified as a virulent factor of CSFV.

Importantly, the identification of these genetic determinants of virulence have been instrumental for providing researchers with a rational approach for designing an improved marker CSF LAV. Conceivably, the genetic introduction of mutations containing one or several of these determinants into a CSFV cDNA infectious clone may lead to the creation of a safer and more efficient vaccine with increased utility and ultimately producing a novel vaccine with

DIVA capabilities which will be discussed in more detail in the next section.

CP7_E2alf Marker Vaccine Candidate

In 2009 an international collaboration composed of 17 multidisciplinary scientific partners from 11 different countries was established to complete a project called

CSFV_GODIVA. The project was initiated because despite extensive efforts made to eradicate

CSFV from the European Union (EU), CSFV continues to be a problem in several countries where the wild boar population are endemically infected with CSFV. These wild boar serve as a continual threat for introduction of CSFV into the domestic pig population. Due to the high costs of the large scale culling and the ethical objections to vast killing of healthy pigs associated with the current “stamping out” control policy and restrictions on prophylactic vaccination usage, there has been an ongoing debate over the possible use of emergency vaccination upon development of an approved CSFV DIVA vaccine within the EU. Unfortunately, the C strain vaccine which has historically and effectively been used for vaccination against CSFV does not possess the ability for serologically differentiating vaccinated animals from animals infected with CSFV field isolates. As a result, in order to ease trade restrictions for domestic pigs, potent marker vaccines are required that allow for reliable differentiation of CSFV infected from vaccinated animals. Although the DIVA E2 subunit vaccines have the potential for differentiation they lack some critical properties that live vaccines offer such as fast and sound protection after single application, prevention of trans-placental infection, and suitability for use via the oral application route. Moreover, the CSF E2 subunit vaccines are no longer commercially available due to the overall lack of specificity exhibited by the current discriminatory ELISA assays based on detection of Erns in the marketplace. Resultantly, the

CSFV_GODIVA project was therefore dedicated to improving the tools and strategies for the prevention and control of CSF.

The principal objective of CSFV_GODIVA project was to develop a marketable live CSF marker vaccine candidate that could be intramuscularly administered to domestic pigs and orally administered via baits to wild boar. Although several experimental CSFV marker vaccines have been developed, none of these candidates at the time of the project’s conception were close to market authorization. It was therefore of utmost importance to select candidates to bring them to the level of licensing. To this end, two promising chimeric pestivirus marker vaccine candidates,

“flc11” (van Gennip et al., 2000) and “CP7_E2alf” (Reimann et al., 2004) were selected for further development. The flc11 chimeric mutant virus contained the genetic backbone of CSFV

C strain in which the CSFV ERNS was exchanged for the homologous gene of BVDV II 5250 while the CP7_E2alf recombinant virus contained the E2 sequence of CSFV strain Alfort 187 in the genetic background of BVDV strain CP7. Comparative animal trials were conducted by two independent and separate research groups for both vaccine candidates via intramuscular and oral administration. The performance of the marker vaccine candidates in vaccination/challenge experiments were compared to efficacy of the C-strain vaccine in order to choose a final candidate for proceeding with registration and licensing. In addition, immunity induction to vaccination was assessed by intranasal challenge with highly virulent CSFV strain Koslov at 7,

14, and 21 days post vaccination. Both vaccines proved to be safe and yielded full protection upon challenge 21 days post vaccination. In addition, the CP7_E2alf mutant virus was also able to mediate protection against lethal challenge at 14 days post vaccination. After reviewing the data collected from these independent comparison trials, it was determined that future efforts

would be directed towards improving mutant virus CP7_E2alf for acquiring licensure as a CSFV live marker vaccine.

A Master Seed stock was produced for the CP7_E2alf vaccine candidate and evaluated in animals. The main objective of this study was to assure the scaled up Master Seed stock of

CP7_E2alf vaccine retained its efficacy under laboratory and field conditions in naïve piglets

(http://cordis.europa.eu/result/rcn/53691_en.html). CP7_E2alf vaccine candidate was tested in 6-

10 week old piglets by intramuscular and oral administration. All piglets were challenged with virulent CSFV Koslov strain 4 weeks after vaccination. Resultantly, the vaccine proved to be effective against lethal CSFV challenge with the exception of one orally vaccinated piglet (out of

15) which succumbed at 7 days post-challenge. In addition, immunization and challenge trials showed that the antibody titers after a single intramuscular or oral vaccination were stable for a minimum of 6 month and fully protected from lethal CSFV challenge. Subsequently, the impact of maternal immunity and effect on pregnant animals was analyzed. CP7_E2alf vaccine provided protection against severe clinical signs and mortality in piglets with maternally derived antibodies (MDA) against CSF C-strain. The DIVA potential of the vaccine in these piglets was proven as well. However, when the experiment was repeated in piglets with MDA against CSF induced by CP7_E2alf vaccination in pregnant sows, orally vaccinated piglets displayed only partial protection. Lastly, the vaccine showed a reduction but not full prevention of transplacental infection. In all, the vaccine was deemed to be safe and efficacious after immunization of piglets and pregnant sows.

Notably, most of the CSF outbreaks in domestic pigs in Europe are a result of outbreaks in the wild boar population. For this reason, CP7_E2alf marker vaccine was tested for efficacy of oral vaccination in wild boar and shown to induce similar protection as the classical C-strain vaccine in wild boar (Koenig, 2007). The vaccine was subsequently evaluated as a bait vaccine for wild boar in field trials in France and Italy. Virus neutralizing antibodies were detected in

36.3% of the wild boars tested after a single vaccination campaign. However, due to the absence of a positive marker in the CP7_E2alf vaccine formulated baits it was not possible to confirm whether analyzed animals had actually consumed the baits. The main obstacle for bait uptake appears to be the availability of other food sources. As such, several new baiting systems were tested, unfortunately, these systems failed to enhance the uptake by young boars. Overall, it was determined for effective control and eradication of CSF in the wild boar population to occur; three vaccination campaigns should be performed per year for a minimum of 6 consecutive years to effectively eradicate the disease.

Importantly, DIVA vaccines are only useful when accompanying DIVA diagnostics are available. As such, two commercially available Erns-based discriminatory ELISAs were evaluated for use with CP7_E2alf vaccine. Both tests were found to be only suitable for application on a herd basis. In particular, the specificity of the tests was too low to reliably discriminate vaccinated from CSFV infected pigs. In successive studies it was further shown that one of the tests is not specific for CSFV, as pigs infected with other members of the pestivirus family also reacted positively for Erns antibodies (Schroeder et al., 2012). The PrioCHECK

CSFV Erns ELISA proved to give the best results against the CP7_E2alf vaccine candidate.

Despite several attempts to optimize the assay’s performance, the test still exhibited deficiencies

in both, sensitivity and specificity, as well as problems with robustness and reproducibility

(Blome et al., 2012). Resultantly, improved DIVA ELISAs have been developed under CSFV

GO_DIVA project to be used in conjunction with CP7_E2alf vaccine candidate and are pending validation for manufacturing at the present time.

Lastly, the usefulness of an emergency vaccine largely depends on how early the vaccine is able to provide protection against lethal infection. To this end, several protection studies have been performed for Cp7_E2alf vaccine candidate. As stated previously, during the process of selecting a CSFV vaccine candidate CP7_E2alf mutant virus was shown to protect animals as early as 14 days post challenge. In addition, CP7_E2alf vaccine stock was evaluated in naïve piglets in a set of vaccination/challenge experiments in which groups of animals were intramuscularly vaccinated with CP7_E2alf vaccine and challenged 1 week later with highly virulent CSFV strain Koslov or vaccinated oronasally and challenged 2 weeks after vaccination

(Leifer, I. et al. 2009). In both cases all vaccinated animals were completely protected against lethal CSFV challenge. A subsequent study was performed in which CP7_E2alf vaccine candidate and C strain were compared for efficacy to induce protection at even earlier times post vaccination. Animals were intranasally vaccinated and challenged with moderately virulent

CSFV isolate Bas-Rhin at 2 days post vaccination (Renson et al 2013). After challenge, both groups were only able to provide partial clinical protection against CSFV. Thus, to date the earliest CP7_E2alf recombinant virus is proven to be able provide complete protection against

CSFV is 7 days post vaccination.

The CP7_E2alf vaccine has recently been submitted for registration and its usage will improve future efforts to eradicate CSF in Europe. From simulation models of EU CSF spread, it has been concluded that emergency vaccination with this type of DIVA vaccine is a better control alternative than the present policy of stamping out and will prevent mass-destruction of non-infected livestock when accompanied with rapid testing procedures. This method will allow for the demonstration of absence of virus and ultimately expedite the time frame in order to regain CSFV free status.

Flag T4v Marker Vaccine Candidate

Our research group has also successfully developed a putative CSFV marker vaccine called Flag T4v; however, our vaccine differs from the European CP7_E2alf vaccine candidate as it contains a negative and positive antigenic marker (Fig. 6) (Holinka et al. 2009). This marker vaccine was produced by introducing two separate genetic determinants of viral virulence into the backbone of CSFV strain Brescia (Risatti et al., 2005b; Risatti et al., 2006). The first of these determinants was previously discovered by the random insertion of a transposon into the carboxyl end of glycoprotein E1 at base pair 2422 in the virulent CSFV Brescia backbone.

Disruption of this region produced an attenuated phenotype in animals. We further modified the

19mer transposon by altering the insertion sequence to include the nucleotide sequence of a well characterized synthetic epitope, Flag® (Sigma, St. Louis, MO) (Fig. 4). This inserted Flag epitope was designed to serve as a positive antigenic marker. In addition, Flag T4v also contains a negative antigenic marker resulting from modifications in a region of E2 glycoprotein known to interact with monoclonal antibody WH303 (mAbWH303) which has a specificity for CSFV

(Edwards et al., 1991). This highly conserved CSFV epitope was mutated to contain the corresponding homologous amino acid sequence of BVDV (Risatti et al., 2006). Shown in Fig. 5 are the progressive mutations which were introduced in this region to determine the minimum alterations required to eliminate total reactivity with mAbWH303. By plaque assay loss of reactivity with mAbWH303 began to be seen between T3v and T4v. As a result of these observations, all further studies were based on mutations required to produce T4v.

Importantly, a critical aspect to be addressed in development of novel live attenuated vaccines for use during a CSFV outbreak in areas free of disease is the ability to produce early protection. Consequently, most CSFV experimental vaccines developed to date have only been tested for their potency against challenge between 2 to 4 weeks post-vaccination. As a result, data is lacking about how efficacious these vaccines are in shorter vaccination-to-exposure trials which would deem them suitable for emergency vaccination. Remarkably, Flag T4v is able to induce a solid protection status in swine from day 3 post-inoculation when administered intranasally, or from day 2 post-inoculation when delivered intramuscularly (Holinka et al.,

2009).

A final essential feature required for a live attenuated CSFV vaccine is the ability to distinguish naturally infected animals from vaccinated animals which necessitates the need for an accompanying diagnostic test. We were able to successfully develop two proof of concept tests to complement Flag T4v. The first set of tests were designed to measure the serologic response against either the Flag epitope or the mAbWH303 epitope which would allow for discrimination between animals immunized with Flag T4v or animals infected with CSFV by enzyme-linked

immunosorbent assays (ELISA). The second set of tests were developed to genetically differentiate Flag T4v from wild-type CSFV based on the annealing of probes to both Flag and

T4 sequences in real-time reverse transcriptase polymerase chain reaction (rtRT-PCR) assays from RNA extracted from infected and vaccinated animal samples (Holinka et al., 2009).

Notably, the latter could prove to be instrumental in monitoring the epidemiological situation during an epidemic as it would provide the ability to test for virus presence even before an efficient antibody response is mounted in CSFV vaccinated or infected animals.

Thesis Research

In light of the DIVA capabilities demonstrated by Flag T4v as well as its remarkable ability to provide complete protection against lethal challenge as early as 3 days post infection, we partnered with a leading animal vaccine manufacturer that had expressed interest in licensing and producing the Flag T4v marker vaccine for commercial use in the field. In collaboration with this company a master seed stock of our virus was produced by the manufacturer at their facility and sent to us for further evaluation. Notably, the master seed stock of Flag T4v was generated by propagation of the virus in a proprietary cell line. This was done in order to yield a single uniform batch of Flag T4v for downstream large scale vaccine production. Consequently, the process to obtain a license for the commercial use of a vaccine is one that is long and rigorous.

One of the initial steps in this procedure is the determination of the safety profile for the master seed stock. This entails the testing of the master seed stock in the host animal to determine the minimum protective dose of the vaccine as well as an overdose study to ascertain whether there are any deleterious side effects associated with the vaccine. In addition, it is required for master

seed stocks derived from live attenuated viruses to demonstrate that they are genetically stable.

This requirement is due to concerns that the master seed stock virus may be transmitted from the host to contact animals potentially causing disease if the viral stock was to retain any residual virulence or if reversion to virulence was to occur. Resultantly, all live vaccines are tested for virulence by virus passage studies, in the host animal, using the most sensitive and usually the youngest animals for which the product is intended.

During the vaccine evaluation process, the master seed stock of Flag T4v was shown to be immunologically potent as well as non-toxic, however, Flag T4v did not sufficiently display genetic stability in reversion to virulence testing when passaged in pigs. Sequence analysis of the revertant virus obtained from the spleen of infected animals showed specific deletions and mutations mainly in the areas encoding for glycoproteins, E1 and E2, when compared with the sequence of the parental Flag T4v. Interestingly, both genetic modifications reside in areas which were previously shown to individually provide the attenuation observed in Flag T4v. Complete deletion of the 18mer insertion at the carboxyl end of the E1 glycoprotein as well as a single amino acid substitution of N850S in the E2 glycoprotein were discovered in the revertant virus,

Flag T4SPv. Surprisingly, the virulent phenotype is only observed when the two mutations occur simultaneously; viruses harboring either mutation exclusively were completely attenuated in pigs. The majority of the non-synonymous mutations observed in Flag T4SPv were restricted mainly to the Flag and T4 regions of the genome. Only two additional non-synonymous mutations, A109V in Npro and A993E in E2, were detected in this virus.

It was hypothesized the genetic stability of Flag T4v could be optimized while still retaining its attenuated phenotype and double antigenic marker elements by altering the codon usage in the Flag and T4 regions. A new viral construct was produced by taking advantage of the redundancy of the genetic code and selecting nucleotide sequences for synonymous codons in these areas that resulted in a construct with a nucleotide sequence differing as much as possible from the initial genomic sequence while maintaining the original amino acid sequence. In addition, a N850G substitution within the T4 region was also introduced due to the observation that reversion of N850S contributes to viral virulence. The resulting virus was called Flag T4Gv and was shown to be stably attenuated when assessed in a reversion to virulence experiment using highly susceptible 5-6 week old piglets. Interestingly, FlagT4Gv could not be detected in tonsil extracts obtained from any of the five successive passage groups. As a result, it was not feasible to ascertain whether the new Flag T4Gv underwent any genomic adaptations as it was passaged in pigs. Consequently, it is not clear how the introduced mutations affecting the codon usage in the Flag and T4 regions increased the genetic stability of Flag T4Gv. Lastly, Flag T4Gv exhibited a similar efficacy to Flag T4v in terms of protecting swine at early and late times post vaccination. These results suggest that Flag T4Gv may be the best suited vaccine candidate developed thus far for emergency usage during a CSFV outbreak.

MATERIALS AND METHODS

Viruses and cells.

The porcine kidney cell line SK6 (Terpstra et al., 1990), free of Bovine Viral Diarrhea

Virus (BVDV) was used throughout this study. SK6 cells were cultured in Dulbecco's minimal essential medium (DMEM) (Gibco, Grand Island, NY) supplemented with 10% gamma- irradiated fetal bovine serum (FBS) (HyClone Laboratories, Logan, UT) and 1% 1 X Antibiotic-

Antimycotic (Gibco). Virulent CSFV Brescia strain (obtained from the Animal and Plant Health

Inspection Service, Plum Island Animal Disease Center) was propagated in SK6 cells.

Minimum Protective Dosage of Flag T4v Master Seed Stock

A master seed stock of Flag T4v was produced by an outside animal vaccine manufacturer and returned to our lab at Plum Island Animal Disease Center for assessment. For determination of minimal protective dose, forty (n=40) 5-6 week old commercial female pigs were used. All animals were acclimated for a period of seven days before being used. Pigs were randomly assigned into four groups of 10 pigs and inoculated intramuscularly in the neck with

5 4 3 10 , 10 , or 10 TCID50 (50% tissue culture infective dose) of the master seed stock of Flag T4v.

The remaining group of 10 pigs was mock vaccinated receiving sterile DMEM media. All

5 animals were intranasally challenged 21 days post vaccination with a lethal dose of 10 TCID50 of highly virulent CSFV Brescia strain, (BICv). Clinical symptoms were scored according to a semiquantitative system based on 10 CSF-relevant criteria described by Mittelhozer et al. (2000).

These 10 CSF parameters are liveliness, body tension, body shape, breathing, walking, skin, conjunctiva, appetite, defecation, and leftover food. Each parameter was judged on a scale of 0 to

3. Accordingly, normal behavior was scored as 0, a slightly altered phenotype was scored as 1, distinct clinical signs were scored as 2, and severe CSF symptoms were scored as a 3. Scores from all 10 criteria were then added up and the total score for each animal was plotted daily. In addition, body temperatures were also recorded daily, with fever defined as values above 104° F.

Blood and serum samples were collected from all animals at 0, 7, 14, and 21 days post vaccination. In addition, blood, serum and nasal swabs were collected from all animals 4, 7, 11,

15 and 21 days post challenge. Lastly, palantine tonsils were removed at 21 days post challenge or at time of death for detection of challenge virus by immunofluorescent histochemistry with mAb 303 (Edwards et al., 1991) and mAb anti-Flag (Sigma-Aldrich, St. Louis, MO).

This animal study as well as all the following studies described here were carried out in accordance with regulations as outlined in the USDA 9CFR-121 and animal welfare act under methodologies described in protocol 171-14-R approved and periodically revised by PIADC

IACUC.

Toxicity and Safety Assessment of Flag T4v Master Seed Stock

The intrinsic safety and toxicity of the master seed stock of Flag T4v was evaluated in seven (n=7) 5-6 week old commercial female pigs. Five pigs were randomly selected to be

7.4 infected intramuscularly with 10 TCID50 of the Flag T4v master seed stock. Pigs were observed over a 21 day period. Rectal temperature as well general clinical symptoms were recorded daily. Presence of local lesions were analyzed by measuring the dimension of the area surrounding the injection site with a caliper daily and comparing with measurements of local reaction of injection site obtained from a group of 2 mock vaccinated animals.

Reversion to Virulence Assessment of Flag T4v Master Seed Stock

Reversion to virulence test was performed using twenty (n=20) 5-6 week old commercial female pigs. All animals were randomly assigned into 5 groups and acclimated for a period of seven days before being used. Reversion to virulence protocol was performed in accordance with

OIE regulations with minor modifications. A group of five animals was intramuscularly

6 inoculated with 10 TCID50 of master seed stock of Flag T4v. Animals were observed for 7 days and clinical signs and body temperature were recorded daily. At day 7 post infection animals were euthanized and palatine tonsils removed. Presence of virus was detected in tonsils by virus isolation evidenced by immunohistochemistry using mAb WH174 for virus detection. Tonsils were then macerated in a 10% w/v suspension and then clarified by centrifugation at 2,000 rpm for 20 minutes. The resulting tonsil tissue supernatants were pooled and used to inoculate a new group of 5 animals. This procedure was repeated for 4 more times for a total number of 5 viral passages in swine.

Viral RNA Extraction from Spleen Sample for Sequencing

QIAGEN RNeasy kit (QIAGEN, Valencia, CA) was used to isolate revertant viral RNA

(Flag T4SPv) from the spleen of one of the affected piglets in passage 5 of the reversion to virulence experiment (Flag T4SPv). The RNeasy protocol used, selectively combines silica- based membrane binding properties with high speed microspin technology. The spleen samples were first lysed and homogenized in a highly denaturing buffer containing guanidine- thiocyanate. Subsequently, 70% Ethanol (Sigma-Aldrich) was added to homogenized sample to provide appropriate binding conditions for RNA to silica membrane. The sample mixture was then applied to RNeasy Mini spin column and centrifuged. Following centrifugation, wash buffer was flowed over spin column to remove contaminants. The bound RNA was eluted off membrane in 20 μl of nuclease free water (Promega, Madison, WI).

Reverse Transcription and PCR amplification of Flag T4SPv for sequencing

The extracted Flag T4SPv RNA was reverse transcribed from RNA to cDNA as follows: a 20 ul reaction mixture [2 U of Moloney murine leukemia virus reverse transcriptase (MMLV)

(Stratagene, Jolla, CA), MMLV 10x buffer, 4 mM deoxynucleosidetriphosphates,(dNTPs), 10 pmol random primers (ThermoFisher Scientific, Waltham, MA), and 5 μl of Flag T4SPv RNA] was incubated 1 h at 37°C. Subsequently, the cDNA of Flag T4SPv was amplified by traditional

PCR, producing 5 overlapping PCR amplicons spanning the entire viral genome. PCR amplification was performed in a 50 μl reaction using Advantage® 2 PCR kit (Clontech, Palo

Alto, CA) [5 μl of 10× Advantage 2 buffer, 400 μM of dNTPs, 10 pmol of each of primer set

(Table 4), 1 μl of Advantage 2 Polymerase Mix and 2 μl of cDNA] and denatured at 95 °C for 1 min followed by 25 cycles at 95 °C for 20 seconds, 52 °C for 20 seconds, and 68 °C for 30 seconds, and a final 68 °C extension cycle for 1 minute on MJ Research Peltier Thermal Cycler

(Bio-Rad). PCR products were resolved on 1% agarose gels stained with ethidium bromide and visualized using gel doc (Bio-Rad, Berkeley, CA). PCR products were purified using QIAGEN

PCR purification kit and sequenced.

DNA Sequencing

Full-length infectious cDNA clones, in vitro rescued viruses, and in vivo rescued virus (Flag

T4SPv) were completely sequenced with 96 CSFV specific primers (Table 2) by the dideoxynucleotide chain-termination method (Sanger et al., 1977). Sequencing reactions were prepared with the BigDye Terminator Cycle Sequencing Kit (Perkin-Elmer, Boston, MA). A

10ul reaction containing BigDye™ Terminator Ready Reaction Mix, 100ng/ul cDNA template and 3.2 µM CSFV primers and cycled on MJ Research Peltier Thermal Cycler (Bio-Rad) as follows: incubate 96 °C for 1 minute, followed by 25 cycles of denaturation at 96 °C for

10 seconds, annealing 50 °C for 5 seconds and extension at 60 °C for 4 minutes. DNA was precipitated and washed with 70% ethanol and resuspended in 10ul of Hi‑ Di™ Formamide.

Reaction products were sequenced on a 3730 xl DNA Analyzer (Applied Biosystems, Foster

City, CA). Resulting Sequence data was assembled and aligned using SequencherTM 5.2.4 software program (Gene Codes Corporation, Ann Arbor, MI). The final DNA consensus sequences presented on average a five-fold redundancy at each base position.

Construction of CSFV chimeric plasmids representing mutations in Flag T4SPv

Sequence analysis of the revertant virus, Flag T4SPv, isolated from the spleen of animals infected with the master seed stock of Flag T4v, revealed the presence of three nonsynomous amino acid substitutions and absence of the Flag epitope insertion residing in the glycoprotein E1 when compared to the sequence of parental Flag T4v genome. The three nonsynomous amino acid mutations were specifically caused by random nucleotide replacements of C687T which resulted in the A109V substitution, nucleotide A2910G resulted in the N850S substitution, and nucleotide C3339A resulted in the A993E substitution (Table 1). To identify the contribution of these mutations on viral virulence observed in Flag T4SPv, a set of recombinant viruses were constructed containing these individual nonsynomous amino acid mutations in both the presence and absence of the Flag epitope insertion. As a result, three mutant plasmid constructs were developed using a full-length infectious cDNA clone (IC) of FlagT4 (Holinka, et al., 2009) as

DNA template; plasmid Flag T4 699, exchanging nucleotide C at position 699 for T, plasmid

Flag T4 2932, replacing base A at position 2932 for G, and plasmid Flag T4 3351, substituting nucleotide C at position 3351 for A. These same mutations were also introduced into the full- length infectious cDNA clone of T4 (Risatti, et al., 2006), a DNA template containing the T4 mutation but lacking the Flag insertion. The plasmids using the T4 template were designated T4

699, T4 2932 and T4 3351 respectively. All 6 plasmids were generated using PCR-based

QuikChange® XL Site-Directed Mutagenesis kit (Stratagene). In order to introduce these nucleotide changes into either the full length plasmid of FlagT4 or the plasmid of T4, a set of mutagenic oligonucleotide primers were designed in accordance with manufacturer’s guidelines

(Table 3): primer length between 25 and 45 nucleotides, melting temperature of ≥78 °C and a

GC content of 40% with desired mutation centered within probe with ~ 10-15 nucleotides of correct sequence on either side (Table 4). Each mutagenic reaction contained ~100ng of dsDNA template and 125ng of the appropriate forward and reverse primer set. Reactions were thermal cycled according to manufacturer’s suggested cycling parameters in order to synthesize cDNA mutant strands. Following mutagenesis, the reactions were treated with 10 U of the endonuclease

Dpn1 to eliminate methylated nonmutated parental DNA templates, and extended cDNA products were transformed into XL Gold ultra competent E. coli cells (Stratagene) pretreated with β-mercaptoethanol (Stratagene) for increased transformation efficiency and plated on terrific broth agar plates with 100ug/ml of Carbencillin (Teknova, Hollister, CA). Plates were incubated overnight at 37°C. The following day individual colonies were picked and again grown overnight in TEKgro terrific broth with 100ug/ml of Carbencillin at 37°C (Teknova). All plasmid

DNAs were purified using QIAGEN highspeed maxi kit. The QIAGEN plasmid purification protocol is based on a modified alkaline lysis procedure. The bacteria were first lyzed under alkaline conditions and then the lysates were subsequently neutralized. Bacterial lysate were next cleared without centrifugation using QIAfilter cartridges. After lysate clearing, the samples were purified on QIAGEN-tips packed with QIAGEN proprietary anion-exchange resin. Plasmid

DNAs bind to QIAGEN resin under the low-salt buffer conditions. Contaminants, such as, RNA, proteins, dyes, and low molecular weight impurities were removed with a medium-salt buffer wash. Plasmid DNAs were eluted in a high-salt buffer and then concentrated and desalted by isopropanol (Sigma-Aldrich) precipitation. Finally, the purified full-length cDNA plasmids were sequenced in their entirety to verify the presence of the desired mutations and absence of undesired random mutations as previously described.

Construction of CSFV Flag T4Gv

The full-length DNA plasmid of Flag T4 was used as DNA template in which mutations were introduced to alter codon usage in order to produce a new plasmid called Flag T4G.

Mutations were introduced by site-directed mutagenesis using QuikChange® XL Site-Directed

Mutagenesis kit as described above. In order to introduce the altered codon usage in Flag and T4 regions (Fig. 8) a series of 4 mutagenic oligonucleotide primer sets were designed in accordance with manufacturer’s guidelines, as described above (Table 7). Notably, 3 sets of primers had to be designed to introduce alterations to the Flag insertion codon usage due to limitations of the site directed mutagenesis protocol in which the efficiency of the reaction significantly decreases after a maximum of 6 base pair substitutions (Fig. 8). Each primer set was individually used in the site directed mutagenesis protocol to introduce mutations progressively into Flag T4 plasmid

DNA. After each site directed mutagenesis reaction, the resulting plasmid DNA was transformed into E. coli and then isolated from bacteria using the QIAGEN plasmid purification protocol. The new plasmid DNA from each reaction was then used as the template to add the next set of mutations through site directed mutagenesis. This process was repeated a total of four times in order to produce the Flag T4G cDNA construct. Lastly, the Flag T4G plasmid was sequenced in its entirety to verify the presence of the desired mutations and absence of any undesired random mutations as described previously.

Rescue of CSFV chimeric viruses

Full-length genomic clones were linearized with SrfI and in vitro transcribed using the T7

Megascript® system (Ambion, Austin, TX), and RNA precipitated with LiCl. RNA was

transfected into SK6 cells by electroporation at 500 volts, 720 ohms, 100 watts with a BTX 630 electroporator (BTX, San Diego, CA). Cells were plated in 6 well tissue culture plates (Corning

Life Science, Corning, NY) and incubated for 4 days at 37C and 5% CO2. In order to detect viral infectivity, one well of each plate was fixed and stained by immunochemistry. Fixation was carried out using a 50% (vol/vol) ethanol-acetone (Fisher Scientific, Pittsburg, PA) solution and incubated for 30 minutes at room temperature and subsequently blocked with DMEM (Gibco) supplemented with 2 % normal horse serum (Vector Laboratories, Burlingames, CA) to prevent non-specific binding. Immunohistochemistry staining was performed using the CSFV E2 specific monoclonal antibody (mAb) WH303 or mAb174 in combination with the Vectastain

ABC® kit (Vector Laboratories) (Risatti et al., 2003). Cells and supernatant were harvested from remaining wells for stock virus using rubber cell scrapers (Corning Life Sciences, Corning, NY).

Stocks of rescued viruses were stored at -70ºC.

In Vitro and In Vivo Rescued Flag T4G Viral RNA and NGS Sequencing

Viral RNA was isolated from in vitro produced viral stock of Flag T4Gv and tonsil tissues from Flag T4Gv infected pigs using TRIzol® LS Reagent method (Life Technologies,

Grand Island, NY, USA). 3 Parts Trizol LS was added to 1 part tissue sample/virus stock for efficient lysis. Chloroform (J. T. Baker, Center Valley, PA) was added to sample and subsequently centrifuged. Centrifugation resulted in the reaction mixture separating into 3 phases, with the upper aqueous phase containing the RNA. Next, the upper layer was transferred to a clean nuclease-free polypropylene microcentrifuge tube and the RNA was precipitated from

solution with isopropanol (Sigma-Aldrich). The precipitated RNA was washed with 70% ethanol

(Sigma-Aldrich) to remove impurities, and then resuspended in nuclease free water (Promega).

The RNA concentration of in vitro rescued Flag T4Gv, and Flag T4Gv tonsil samples and were determined using the Qubit® RNA high-sensitivity (HS) assay kit (Life Technologies) and read on a Qubit 2 fluorometer (Life Technologies). One microgram of viral RNA was enzymatically fragmented to obtain blunt-end fragments in a length range of 100 to 300 bp using the enzyme RNase III (Life Technologies) and incubated at 37°C in a MJ Research Peltier

Thermal Cycler (Bio-Rad). Ion adapters were hybridized to fragmented RNA, hybridized fragmented RNA was reverse transcribed to cDNA and cDNA was ligated to ion library barcodes and amplified using the Ion total RNA-seq v2 kit (Life Technologies). 1ul of amplified cDNA was loaded onto Agilent DNA 1000 chip (Agilent, Santa Clara, CA, USA) and quantified using 2100 bioanalyzer (Agilent). Using the 2100 expert software, a smear analysis was performed to quantify the molar concentration (nM) of the cDNA libraries in order to determine the library dilution required for template preparation. After dilution of cDNA library to a concentration of 20pM was performed, the cDNA library was clonally amplified onto Ion Sphere particles (ISPs), generating template-positive ISPs using the Ion PGM template OneTouch 2 200 kit (Life Technologies) with the Ion OneTouch 2 instrument (Life Technologies). Before proceeding to enrichment, quality assessment of nonenriched template-positive ISPs was performed using the Ion Sphere quality control (QC) assay kit (Life Technologies) and a Qubit 2 fluorometer. The template-positive ISPs were then enriched using the Ion PGM template

OneTouch 2 200 kit (Life Technologies) and Ion OneTouch enrichment system (ES) instrument

(Life Technologies) to eliminate template-negative ISPs and to denature cDNA on template- positive ISPs. Using the Ion PGM 200 sequencing v2 kit (Life Technologies), enriched template

ISPs were prepared for sequencing, loaded onto either an Ion 316 chip v2 (Life Technologies), and run on the Ion PGM sequencer (Life Technologies). The sequences obtained were then trimmed using Galaxy (https://usegalaxy.org) NGS QC and manipulation tools. Sequences were aligned and analyzed using Sequencher 5.2.2 (Genecodes) and CLC Genomics Workbench

(CLCBio) software.

Protection of Flag T4Gv Animal Study

Protection experiments were performed using commercial female pigs weighting 40-60 lbs. Animals were randomly allocated in three groups harboring five (n=5) animals each. Two

5 groups were intra muscularly immunized with 10 TCID50 of Flag T4Gv. The third group was mock vaccinated with sterile DMEM media. One Flag T4Gv vaccinated group of animals were

5 intranasally challenged at 7 days post vaccination with 10 TCID50 BICv. The remaining two

5 groups were challenged at 21 days post immunization with 10 TCID50 BICv. Clinical signs and body temperature were recorded daily throughout the experiment as previously described.

Viremia was determined for samples collected at 4, 7, 11, 15, and 21 days post-challenge by virus titration in SK6 cells as described below.

Reversion to Virulence Assessment of Flag T4Gv

Reversion to virulence assessment for Flag T4Gv was implemented using twenty (n=20)

5-6 week old commercial female pigs. Animals were randomly assigned into 5 groups and acclimated for a period of seven days before beginning experiment. Reversion to virulence protocol was performed as described previously. The initial group of five animals were intra muscularly inoculated with 106 TCID50 Flag T4Gv produced in our lab. Clinical signs and body temperatures were recorded for a period of 7 days. Seven days post infection all five animals were euthanized and palatine tonsils collected. Virus isolation was performed to determine presence of virus in tonsils and confirmed by immunohistochemistry using mAb WH174 for virus detection. Tonsils were then macerated in a 10% w/v suspension and a pool of tonsil tissues suspension and used to inoculate a new group of 5 animals. This process was repeated a total of four intervals resulting in 5 passages of Flag T4Gv through animals.

Early Protection of Flag T4Gv Animal Study

To determine the earliest onset of FlagT4v protection swine were allotted into 6 groups of

5 animals each. Five groups were intramuscularly (IM) inoculated with 105 TCID50 of Flag

T4Gv, and one additional group was mock inoculated. All groups were then challenged intranasally with 105 TCID50 of virulent BICv at day 0, 1, 2, 3, 5 or 7 post Flag T4Gv inoculation. Clinical evaluation and sampling of animals was performed as described as earlier.

Virus Titrations

End point titrations for virus stocks and clinical samples were performed by preparing ten-fold dilutions in a diluent of DMEM with 1% antibiotics. The serial dilutions were transferred into six wells each of 96 well tissue culture plates (Corning Life Sciences, Corning,

NY) seeded with 1.0 X 104 cells/ml of SK6 cells. After 4 days of incubation at 37C and 5%

CO2, cells were fixed with a 50% (vol/vol) ethanol-acetone (Fisher Scientific, Pittsburgh, PA) solution for 30 min. at room temperature and subsequently blocked with DMEM supplemented with 2 % normal horse serum (Vector Laboratories, Burlingames, CA) to prevent non-specific binding. Immunohistochemistry staining was performed using either the CSFV E2 specific mAb

WH303, (Edwards et al., 1991) in combination with the Vectastain ABC® kit (Vector

Laboratories Burlingames, CA) (Risatti et al., 2003) in order to detect viral infectivity. The monolayers were read microscopically for stained cells (Wensvoort et al., 1986). Titers were calculated using the method of Reed and Muench (Reed et al., 1938) and expressed as log 10

TCID50/ml. As performed, test sensitivity was > 1.8 TCID50/ml.

Virus replication kinetics

Characterization of the replication kinetics of the Flag T4Gv was compared to parental virus, Flag T4v and WT Bresciav by the performance of multistep growth curves. Six well tissue culture plates seeded with 1.0 X 104 cells/ml of SK6 cells were infected at a multiplicity of infection (MOI) of 0.01 TCID per cell and allowed to adsorb for 1 hr at 37C and 5% CO2. The virus inoculum was removed from the cells, and the cells were washed and incubated in freshly

prepared media of DMEM containing 10% уIR FBS and 1% antibiotics at 37C and 5% CO2. At the following hours post infection; 2, 6, 24, 48 and 72 the respective tissue cultures plates were put at - 70° C to stop virus replication. Virus yield was determined by end point titration in SK6 cells.

Plaque Assays

Six well tissue culture were seeded with 1.0 X 104 cells/ml of SK6 cells and incubated at

37°C and 5% CO2 for 24h. After 24h, each well was independently infected with either Flag

T4Gv or BICv (50 to100 TCID) and subsequently overlaid with 0.5% agarose and incubated for

4 days at 37°C and 5% CO2. Plates were fixed and stained by immunohistochemistry as previously described.

Early Protection Animal Study for Assessment of Cytokine Expression

In an effort to elucidate probable mechanisms mediating the early protective immune response observed in animals infected with Flag T4v and challenged at 3 days post infection with virulent BICv an early innate immune response study was conducted using 11 animals. Three

5 groups of 3 pigs each were intramuscularly infected with either 10 TCID50 of FlagT4Gv and

5 euthanized at 3 days post infection, or infected with 10 TCID50 BICv and euthanized 3 days later

5 or infected with 10 TCID50 of FlagT4Gv and intranasally challenged 3 days post infection and euthanized 3 days post challenge. An additional two animals were used as naive controls. Tonsils

and draining lymph nodes were collected after euthanasia from all 11 swine. Blood samples were also collected at time of death.

Quantification of Cytokine Expression in Serum

Sixteen commercially available ELISA kits were used to measure presence and quantify cytokine expression following manufacturer protocols (MyBioSource, San Diego, CA), were used for detection and quantification of various cytokines in serum of animals euthanized 3 days

5 5 after either being intramuscularly infected with either 10 TCID50 of FlagT4Gv, 10 TCID50

5 5 BICv or 10 TCID50 of FlagT4Gv/challenge 3dpi with 10 TCID50 BICv or mock infected animal. Concentrations of cytokines in serum were calculated by extrapolation in a standard curve simultaneously determined with recombinant porcine IFNα (PBL Assay Science).

MyBioSource ELISA kits: Interleukin 8 (IL-8), Interleukin 1 alpha, Interleukin 1beta, Interleukin

2 (IL-2), Interleukin 5 (IL-5), Interleukin 6 (IL6), Anti-Interleukin 10, Interleukin 12 p35,

Interferon-induced GTP-binding protein Mx1 (MX1), Interferon beta, IFN-beta/IFNB, Monocyte

Chemotactic Protein 2 (MCP2), Protein Kinase R (PKR), Transforming growth factor beta1

(TGF-beta1), Total Tumor Necrosis Factor, VCAM-1/CD106, 2',5'-Oligoadenylate Synthetase 1.

RNA extraction and cDNA synthesis for cytokine assay

Total cellular RNA was extracted from tonsils of swine euthanized 3 days after either

5 5 5 being intramuscularly infected with either 10 TCID50 of FlagT4Gv, 10 TCID50 BICv, 10

5 TCID50 FlagT4Gv/challenge 3dpi with 10 TCID50 BICv or mock infected animals. RNA was extracted using a RNeasy Mini Kit (Qiagen) following the manufacturer’s protocol. Contaminant

DNA was removed by DNAse treatment using Turbo DNA-free (Ambion, Austin, TX). After

DNAse treatment, genomic DNA contamination of RNA stocks was assessed by real-time PCR amplification targeting the porcine β-actin gene. Total RNA was quantified using a NanoDrop

1000 (Thermo Scientific, Waltham, MA). cDNA was synthesized with a High Capacity cDNA

Reverse Transcription Kit (Applied Biosystems, Foster City, CA) using random hexamer primers

(Invitrogen, Carlsbad, CA) in a 100 μl reaction containing 1500 ng of total RNA.

Quantitative reverse transcription real-time polymerase chain reaction (qRT-PCR)

For gene expression quantification, first-strand cDNA was amplified by real-time PCR using Power SYBR® Green PCR Master Mix (Applied Biosystems, Foster City, CA) with primer pair sets illustrated in Table 10. A 50 μl reaction contained 25 μl of Power SYBR® Green PCR

Master Mix, 5 μL of cDNA, and 400 nmol/μl of each primer. Cycling conditions were as follows: activation of the AmpliTaq Gold® Polymerase at 95 °C for 10 min, followed by 40 cycles of denaturation at 95 °C for 15 seconds, and annealing and extension at 60 °C for

1 minute, with a melting curve analysis performed at the conclusion of the PCR assay. Real-time

PCRs were run with an ABI 7900HT real-time PCR system (Applied Biosystems).

Quantification of gene expression was performed by the 2−ΔΔCt method (described in ABI

PRISM® 7700 Sequence Detection System User Bulletin #2 (PN 4303859)), and was analyzed with ABI software: RQ Manager 1.2.

RESULTS

Minimum Protective Dosage of Flag T4v Master Seed Stock

In collaboration with a major animal vaccine company a master seed stock of Flag T4v was produced by the manufacturer at their facility and sent to us for further evaluation. Notably, the master seed stock of Flag T4v was generated by propagation of the virus in a proprietary cell line. This was done in order to yield a single uniform batch of Flag T4v for downstream large scale vaccine production. Consequently, potential vaccines must undergo a rigorous licensing process to ensure the safety, efficacy, purity and potency before it can be approved for marketing. One of the initial steps in this procedure is the determination of the safety profile for the master seed stock. This entails the testing of the master seed stock in the host animal to determine the minimum protective dose of the vaccine as well as an overdose study to ascertain whether there are any deleterious side effects associated with the vaccine. The minimal protective dose is defined as the dosage that protects at least 80% of the animals in a given study.

Furthermore, determination of the minimum protective dose demonstrates the potency of the vaccine candidate as an immunogen. Importantly, faulty dose selection can result in either selecting a dose that is too low to achieve the desired benefit or one that is too potent, resulting in unacceptable toxicity.

Three groups of 5 animals each were inoculated with dose ranges of 105, 104 and 103

TCID50 of Flag T4v master seed stock. A fourth group of 10 pigs were mock vaccinated. Fever responses and clinical signs were monitored daily following lethal CSFV challenge 21days post

5 infection. Unexpectedly, one animal in the group inoculated with 10 TCID50 of Flag T4v died at day 6 post infection. Importantly, necropsy of this animal showed no evidence of CSFV, ASFV, or FMDV related pathology indicating that death was not due to inoculation with Flag T4v or a result of unintentional introduction of another foreign animal disease used at our facility. All remaining nine animals in this group survived lethal challenge remaining clinically normal with a few pigs occasionally exhibiting body temperatures slightly above 104°F (Table 5 and Fig.

7A). In contrast, two animals developed fever 6 days post challenge in the group inoculated with

4 10 TCID50 Flag T4v (Table 5 and Fig. 7B). These two animals were euthanized at days 9 and 15 post challenge. At necropsy, pathological findings indicative of CSFV were found in both

3 animals. Similarly, two pigs in the group infected with 10 TCID50 Flag T4v developed fever at days 3 and 5 post challenge (Table 5 and Fig. 7C). These pigs were euthanized at days 8 and 13 post challenge presenting CSFV related symptoms/lesions. The remaining animals in all 3 Flag

T4v infected groups remained clinically normal. In contrast, all animals in the mock vaccinated group presented CSFV related symptoms after challenge and were euthanized at days 7 (two animals), 8 (one animal), 9 (two animals), 10 (two animals) and 11 (three animals) post challenge (Table 5 and Fig. 7D).

Back-titrations of the Flag T4v inoculums revealed that each animal group received approximately one log less of Flag T4v than intended. These findings infer that the minimum

2 3 protective dose of Flag T4v master seed stock is actually 10 TCID50 rather than 10 TCID50.

Remarkably, these results indicate that Flag T4v is a potent immunogen.

In addition, immunohistochemistry and titrations were performed on collected tonsil tissues. For animals infected with a dose of 105 TCID50 of Flag T4v all animals were negative by virus titration and immunohisotchemisty using mAb 303 showing that multiplication and spread of challenge virus is very limited or not present. However, immunohistochemistry with mAb anti-Flag showed the presence of vaccine virus in two animals. These same animals were

4 3 negative in virus titrations. Animals in the groups infected with 10 and 10 TCID50 of Flag T4v no virus was detected using mAb anti-Flag. As expected the two animals in each group that exhibited clinical CSFV symptoms after lethal CSFV challenge were both positive by immunohistochemistry and virus titrations using mAb 303. All mock infected animals were also positive by immunohistochemistry and virus titrations using mAb 303 but negative for presence of vaccine virus as shown by immunohistochemistry with mAb anti-Flag

Flag T4v Safety Test

The intrinsic safety of vaccines should be demonstrated early in the development stage and documented as part of the vaccine licensing process. Five 40-60 lb swine were

7.4 intramuscularly inoculated with 10 TCID50 of the master seed stock of Flag T4v. Animals were observed over a 21 day period. Rectal temperatures, as well as general clinical symptoms were recorded daily. Notably, over the course of the first week post inoculation the injection site of animals vaccinated with Flag T4v was carefully monitored for appearance of local lesions and compared with reactions at inoculation site of the two mock vaccinated pigs. None of the five

animals developed CSFV related symptoms or exhibited adverse reactions at injection sites. The absence of clinical signs of infection or tissue reactions at the application site after vaccination with Flag T4v shows that the candidate vaccine is completely atoxic.

Reversion to Virulence Test for Flag T4v Master Seed Stock.

With live vaccines, there is always a concern that the live attenuated virus might be shed from the host and transmitted to contact animals, causing disease if the virus was to retain residual virulence or revert to virulence. Therefore, all live vaccines are required to be tested for virulence by means of passage studies, in the host animal, using the most sensitive, usually the youngest, animals for which the product is intended. Consequently, putative vaccines are propagated in vivo by inoculating a group of target animals with the master seed stock.

Subsequently, the live attenuated virus is recovered from tissues of these infected animals one week after inoculation and then used to infect the next group of naïve animals. After not less than four virus passages in animals, with at least a total of five groups of animals, the live attenuated virus is isolated from tissue and then fully characterized, using the same procedures used to characterize the master seed stock. This process was used to assess the genetic stability of Flag

T4v master seed stock. Accordingly, the conventional five passage reversion to virulence

5 experiment was performed by intramuscular inoculation of piglets with 10 TCID50 of Flag T4v master seed stock with subsequent passage groups receiving supernatant macerates from pooled virus-positive tonsils obtained from the previous passage. The first three passages of inoculated piglets did not demonstrate clinical signs of CSF. However, during the fourth passage, all animals presented a transient rise in body temperatures, although none of them presented any

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additional CSFV related clinical symptom. In addition, no CSFV related lesions were found at necropsy (data not shown). In the fifth passage two animals were euthanized after exhibiting severe CSFV clinical signs. Necropsy of these animals also revealed presence of CSF related lesions. The other three animals presented transient fever between days 4 and 5 accompanied by depression, with some minimal CSF related lesions found at necropsy (data not shown). These results indicate that Flag T4v was not able to maintain its attenuated phenotype when successively passed in its natural host.

Genetic Modifications Introduced in Flag T4v During the Reversion to Virulence Protocol

To elucidate the genetic changes in the Flag T4v genome that led to reversion to virulence, full genome sequencing of the virus isolated from the spleen of one of the affected piglets (Flag T4SPv) was performed. The obtained sequence was then compared to attenuated parental Flag T4v. Interestingly, nucleotide sequence comparisons revealed three amino acid substitutions, a deletion of a stretch of 18 amino acids, and one single nucleotide deletion (Table

1). Of the three amino acid substitutions in Flag T4SPv, one mutation was found to affect the structural protein Core. At amino acid position 109, a non-polar hydrophilic alanine was replaced for another non polar amino acid valine. The other two mutations found in Flag T4SPv resided in the glycoprotein E2. The first of these mutations exchanged a polar uncharged hydrophilic amino acid asparagine for a similarly uncharged polar hydrophilic serine at amino acid position 850.

Intriguingly, N850 is located in the center of the T4 mutated region that is associated with the lost of reactivity to mAb WH303 (constituting the negative antigenic marker of Flag T4v) and attenuation of T4v in swine (Risatti et al., 2006). The second amino acid mutation in

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glycoprotein E2 was found at amino acid position 3351 and substitutes a non-polar hydrophilic alanine for a negatively charged glutamic acid. In conjunction with these three amino acid substitutions, Flag T4SPv was found to contain a large deletion of 18 amino acids between residues 688 to 706 in the carboxyl end of glycoprotein E1, which comprises the complete Flag epitope/transposon insertion. Finally, a deletion of a cytosine nucleotide was observed at position

12342 within the 3’UTR of Flag T4SPv (Table 1).

Identification of Flag T4v Mutations Leading to Increased Virulence in Flag T4Sv

In order to identify the individual contribution of each of the mutations found in Flag

T4SPv on reversion to virulence, a set of recombinants viruses were constructed containing either the 3 amino acid substitutions either individually or in combination with deletion of the

Flag insert. Importantly, the amino acid substitutions observed in Flag T4SPv were caused by single nucleotide mutations: nucleotide C699T resulting in the A109V substitution, nucleotide

A2932G resulting in the N850S substitution, and nucleotide C3351A resulting in the A993E substitution (Table 1). Consequently, six different mutant viruses were developed: Flag T4v harboring the A109V mutation (Flag 699v), Flag T4v harboring the N850S mutation (Flag T4

2932v), Flag T4v harboring the A993E mutation (Flag T4 3351v), T4v (lacking the Flag insertion) harboring the A109V mutation (T4 699v), T4v harboring the N850S mutation (T4

2932v), and T4v harboring the A993E mutation (T4 3351v). Each of these recombinant viruses was assessed for virulence in swine. Groups of four 40-60lbs pigs were independently inoculated

4 with 10 TCID50 of the six viruses. The presence of CSF related clinical signs and temperatures were recorded daily throughout the observation period of 21 days. Animals inoculated with these

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recombinant viruses remained clinically normal with the exception of the group of pigs inoculated with T4 2932v which presented clinical disease indistinguishable from that observed in animals infected with virulent CSFV BICv (Table 6). These results suggest that deletion of the

Flag insertion within the E1 glycoprotein, along with the amino acid substitution N850S in the

E2 glycoprotein are responsible for the virulent phenotype observed in revertant Flag T4Sv.

Development of Flag T4Gv

After identifying the genetic changes within Flag T4v that caused the virulent phenotype, in revertant virus Flag T4SPv, the next step was to devise a method to modify the Flag T4v in order to improve its genetic stability while maintaining its attenuation as well as its double antigenic marker elements. In order to accomplish this task we decided to genetically modify the

Flag and T4 areas by taking advantage of degeneracy of the genetic code. It is the degeneracy of the genetic code which accounts for overabundance in the number of codons that encode for the same amino acid, specifically there are a total of 64 codons, three of these codons encode for stop codons while the remaining 61 codons are translated into only 20 different amino acids. The term codon usage is used to describe this phenomenon. Consequently, the genome of Flag T4v was modified in the areas encoding for the Flag and T4 epitopes (amino acid residues at positions 688-706 and 848-852, respectively) by changing the codon usage. By using synonymous variant codons we were able to alter the nucleotide sequences as much as possible from the original Flag T4v nucleotide sequence without changing the protein sequence of Flag

T4v (Fig. 8). In addition, a hydrophilic polar amino acid asparagine at position 850 in the T4 region, was replaced with a non-polar glycine in an effort to help inhibit reversion from

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asparagine to another hydrophilic uncharged polar amino acid, serine, the native residue at this position 850 in virulent CSFV strain Brescia (Fig. 8B). The resulting cDNA infectious clone,

Flag T4G, was obtained by multiple rounds of sequential site directed mutations introduced into full-length infectious cDNA clone of the Flag T4, using primers presented in Table 7 (complete description of all substitutions introduced depicted in Fig.8A and B).

Rescue of Flag T4Gv

Infectious RNA was in vitro transcribed from full-length cDNA infectious clone of the

Flag T4Gv (Fig. 8) and used to transfect SK6 cells. Virus was rescued from transfected cells by day 4 post-transfection. Full-length nucleotide sequence of the rescued virus genome was identical to parental DNA plasmid, confirming that only the predicted mutations were reflected in rescued virus.

In vitro growth characteristics of Flag T4Gv were evaluated relative to wild type CSF

BICv and parental Flag T4v in a multi-step growth curve. SK6 cell cultures were infected at a multiplicity of infection (MOI) of 0.1 TCID50 per cell. Virus was adsorbed for 1 hour (time zero), and samples were collected at 2, 6, 24, 48 and 72 hours post-infection (PI). Samples were titrated in SK6 cells, the presence of virus detected by immunoperoxidase with mAb WH174

(which recognizes an E2 epitope different from that recognized by mAb WH303) and titers using calculated using the Reed and Muench methodology (Reed et al., 1938) and expressed as log 10

TCID50/ml. The results (Fig. 9) showed that Flag T4Gv had an approximately 5-8 fold decrease in the final virus progeny yield relative to wild type CSF BICv and 5-8 times more progeny

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relative to parental Flag T4v (reaching average titer values of 7.25, SD=0.09; 6.3, SD=0.17; and

7.62, SD=0.09 TCID50/ml, for Flag T4Gv, Flag T4v, and BICv, respectively).

The antigenic profile of Flag T4Gv was evaluated by assessing the reactivity of the virus against mAbs anti-Flag, WH303 and WH174 and comparing the antigenic profile with that of the control virus, BICv. Flag T4Gv strongly reacted with mAbs anti-Flag and WH174, but did not react with mAb WH303 (directed against the highly conserved epitope in the T4 region) (Fig.

10). Wild-type CSFV BICv reacted with mAbs WH174 and WH303 but lacked reactivity against mAb anti-Flag. Thus, Flag T4Gv displayed the expected antigenic reactivity profile.

Assessment of genetic stability of Flag T4Gv Using the Reversion to Virulence Protocol

Flag T4Gv genetic stability was then assessed using the reversion to virulence protocol set by the OIE under the same conditions used to evaluate Flag T4v. A group of five, 5-6 week

6 old female piglets were intramuscularly inoculated with 10 TCID50 of Flag T4Gv and euthanized 7 days later. Palatine tonsils from each piglet were removed and macerated in DMEM to obtain a 10% weight/volume suspension which was then used to intramuscularly inoculate another group of naïve female piglets. This process of inoculation-euthanasia-inoculation was repeated a total of five times. In all cases animals were kept under observation during a 7 day period and clinical signs and body temperature were recorded daily. At day 7 post infection animals were euthanized and their palatine tonsils removed. Detection for the presence of the

Flag T4Gv was tested by infecting SK6 cell cultures with each passage of the tonsil extracts and evidence by immunohistochemistry using mAb WH174. None of the animals in any of the

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passages exhibited CSFV related symptoms. Moreover, presence of Flag T4Gv was only

3.2 detected at low titers (10 TCID50/ml) in tonsil extracts obtained from the initial group of animals inoculated. All subsequent tonsil extracts were negative by virus isolation. In contrast to

Flag T4v, these results demonstrate that Flag T4Gv is genetically stable and able to maintain an attenuated phenotype under direct pressure for reversion to virulence.

Flag T4Gv Protects Pigs Against Lethal CSFV Challenge

The ability of Flag T4Gv to induce protection against virulent BICv was assessed in early and late vaccination-exposure experiments. Two groups of pigs (n=4) were intramuscularly inoculated with 105 TCID50 Flag T4Gv and challenged at either 7 or 28 days post infection with

105 TCID50 of virulent CSF BICv. Mock-vaccinated control pigs receiving BICv only developed anorexia, depression, and fever by 4 days post-challenge and died or were euthanized in extremis

9 to 10 days post challenge (Table 8). Notably, Flag T4Gv induced complete clinical protection at 7 days and 28 days after immunization. All pigs survived infection and remained clinically normal, without fever (Table 8). Viremia in vaccinated and challenged animals was examined at

4, 6, 8, 11, 14 and 21 DPC. Detection was performed using mAb WH303, which reacts specifically with the challenge virus BICv. As expected, in mock-vaccinated control animals, viremia was observed within 4 days after challenge, with virus titers remaining high until the last time point tested before animals died or were euthanized (Fig. 11). Conversely, animals inoculated with Flag T4Gv and challenged with BICv at 7 or 28 DPI did not present any detectable viremia (Fig. 11). In addition, the presence of BICv could not be detected in blood

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samples during the 21day observation period after challenge. These results indicate that protection induced by Flag T4Gv was complete, preventing both the presentation of CSF-related clinical signs and the replication of the challenge virus.

Assessment of Early Protection Induced by Flag T4Gv

To assess the efficacy of Flag T4Gv as an experimental vaccine in terms of inducing early protection against challenge with the virulent CSFV BICv. Five groups of animals (n=5)

5 were intramuscularly inoculated with 10 TCID50 of Flag T4Gv and were challenged by the

5 intranasal administration of 10 TCID50 of BICv 1, 2, 3, 5, and 7 days later. Clinical signs and body temperatures were recorded daily over the course of the 21 day observation period. Mock infected animals presented a typical evolution of the disease starting at 5 days post challenge and evolving to a severe form of the disease with all animals euthanized between days 8 and 10 post challenge. All animals challenged on the first day post Flag T4Gv infection presented disease kinetics similar to that of the mock treated animals with all pigs euthanized between 9 and 11 days post challenge (Table 9). Animals challenged on the second day post Flag T4Gv inoculation presented a heterogeneous phenotypic behavior. Two of the swine developed a severe form of disease starting at 6 days post challenge with disease kinetics similar to mock treated animals with all pigs euthanized between 11 and 14 post challenge. A third animal presented a transient elevation in body temperature without of any additional clinical signs. While the last two pigs in the group remained completely asymptomatic during the 21 day observation period. Importantly, animals in groups challenged on day 3, 5 and 7 after infection with Flag T4Gv also remained

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completely asymptomatic during this same time frame. These results demonstrate that Flag

T4Gv efficiently provides complete protection against the development of CSF disease since 3 days post infection.

Viremia in Animals from Flag T4Gv Early Protection Study

Viremic values in Flag T4Gv infected animals challenge with BICv at 1, 2, 3, 5, and 7 closely followed the appearance of clinical signs and severity of the disease (Fig. 12). Mock treated animals presented typical high titer values at both 4 and 7 days post challenges. Animals challenged by the first day post Flag T4Gv infection also presented high viremia at 7 and 14 days post challenge. While, average viremia titers were significantly lower in the groups of animals challenged at day 2 after Flag T4Gv due to the fact that surviving animals did not present detectable viremia. Viremia in animals challenged at either 3, 5 or 7 days post Flag T4Gv infection were consistently undetectable during the observational time frame. These results indicate that Flag T4Gv can efficient protect animals is able to provide solid protection against lethal challenge in animals after 3 post infection animals became solidly protected against the challenge with the virulent parental virus in terms of both the appearance of clinical signs associated with the disease and replication of the challenge virus.

Detection and Quantification of Cytokine Expression in Early Protection Animal Study

In an effort to elucidate probable mechanisms mediating the early protective immune response observed in animals infected with Flag T4v and challenged at 3 days post infection with

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virulent BICv an early innate immune response study was conducted using 11 animals. Three

5 groups of 3 pigs each were intramuscularly infected with either 10 TCID50 of FlagT4Gv and

5 euthanized at 3 days post infection, or infected with 10 TCID50 BICv and euthanized 3 days later

5 or lastly infected with 10 TCID50 of FlagT4Gv and intranasally challenged 3 days post infection

5 with 10 TCID50 BICv and euthanized 3 days post challenge. An additional two animals were used as naive controls. Tonsils and draining lymph nodes were collected after euthanasia from all

11 swine. Blood samples were also collected at time of death. The levels of multiple cytokines present in serum were measured in each group of animals by ELISA and qRT PCR cytokine assay. Specifically, levels of MCP2, TGF-1, IFN-, IFN-, IFN-, IL1- IL1-, IL2, IL5, IL-6,

IL-8, IL10, IL12-p35, IL12p40, OAS, PKR, TNF, MX-1, and VCAM were assessed using commercial ELISAs following manufacturer protocols (MyBioSource, San Diego, CA). The results of the ELISAs were used to try to establish a correlation between mediators of the innate host immune response and infection with either attenuated Flag T4Gv or virulent BICv. Notably, the levels of most of the detectable cytokines were quite homogeneous among each group (Fig.

13). The average cytokine values did not significantly vary between the virus infected groups nor did those values significantly differ between virus infected pigs and naïve animals. Notably, only

IFN- showed any significant differences between groups. At three days post FlagT4Gv infection, circulating IFN- was determined to have a concentration of 536.72 ng/ml

(SD+36.26). This concentration was much higher than those found in animals infected BICv which was 174.12 ng/ml (SD+45.85) or for swine infected with FlagT4G/BICv challenged which was 14.4 ng/ml (SD+10.37). Similarly, naïve animals presented values of 8.19 ng/ml

(SD+1.34) for IFN-. These results suggest that IFN- may play a significant role as a mediator of innate immune response against BICv challenge.

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DISCUSSION

The majority of CSF control programs in enzootic areas are based on prophylactic vaccination of domestic pigs. Vaccination with live attenuated vaccine (LAV) strains induce protective immunity in pigs within a few days after vaccination, even before neutralizing antibodies can be detected, providing lifelong immunity against disease (Biront and Leunen,

1988; Terpstra, 1991). However, current LAVs lack DIVA capabilities that would allow differentiation between infected and vaccinated animals, limiting their use as an emergency control strategy in the event of an outbreak in CSFV-free regions. CSFV subunit marker vaccines with DIVA capabilities have been developed using recombinant E2 envelope protein expressed in baculovirus system (Hulst et al., 1993; van Rijn et al., 1996; 1999). These subunit vaccines were commercially available but have since been removed from the market place due to insufficient sensitivity and specificity of their accompanying ELISA assays. In addition, these subunit vaccines were not ideal for emergency vaccination against CSFV due to delayed onset of protective immunity which does not fully develop until nearly two weeks after initial vaccination

(Bouma et al., 2000; Uttenthal et al., 2001). Resultantly, there is an increased need for developing LAVs against CSFV with DIVA capabilities (Beer et al., 2007; Deng and Chen,

2007). These vaccines, while inducing a rapid and effective immunity, should harbor antigenic markers for serological distinction of infected from vaccinated animals. In addition, a rapid molecular diagnostic test, such as real-time RT-PCR, should also be constructed for early detection and differentiation between vaccinated and wild-type infected field isolates.

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This research project describes the development of a novel CSFV live attenuated vaccine candidate, Flag T4Gv, based on the genetic alteration of codon usage in a previously described double marker vaccine virus, Flag T4v (Holinka et al., 2009). Flag T4v was a rationally designed

LAV that was developed via genetic modification of virulent BICv, a virus derived from a full- length cDNA copy of CSFV Brescia strain (Risatti et al., 2005a). The double antigenic marker

LAV Flag T4v was obtained by combining two genetic determinants of attenuation: a) insertion of 19 amino acid residues in the carboxyl end of E1 (Risatti et al., 2005b), and b) mutation of the mAb WH303 epitope in E2 glycoprotein (Risatti et al., 2006). The 19mer amino acid insertion was further modified to include a synthetic Flag® epitope (Holinka et al., 2009). Intranasal or intramuscular administration of Flag T4v protected swine against virulent CSFV Brescia strain at early (2 or 3 days) and late (28 days) times post-inoculation. Flag T4v induced antibody responses in pigs that reacted strongly against the Flag epitope but failed to inhibit binding of mAb WH303 to a synthetic peptide representing the WH303 epitope (Holinka et al., 2009).

When evaluating the genetic stability of Flag T4v via serial passages in highly susceptible

5-6 week old piglets, reversion to virulence was observed in animals in the 5th and final passage group. The revertant virulent virus was rescued from the spleen of one of the sick piglets.

Nucleotide sequencing revealed three amino acid substitutions, a deletion of a stretch of 19 amino acids, and one single nucleotide deletion. A further study aimed at determining the contribution of these mutations on reversion to virulence revealed that deletion of the 19mer Flag insertion in combination with a single amino acid substitution at position N850S were required for producing the virulent phenotype. Notably, viruses harboring either mutation individually were completely attenuated in pigs. Interestingly, the majority of the non-synonymous mutations

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observed in Flag T4Sv were restricted exclusively to the Flag and T areas of the genome. Only two additional non-synonymous mutations, A109V in Npro and A993E in E2, were observed in this virus. Why these discrete areas of the viral genome are genetically modified after successive passages in piglets remains unclear. Similarly, the occurrence of reversion to virulence has been reported by Tamura et al. (2012) in which a live attenuated CSFV vaccine strain, GPE- virus, was readapted to swine by serial passages in pigs that resulted in reversion to virulent phenotype after the eleventh passage in vivo. The original GPE- strain was obtained through passages in cultured cells and was found to possess 225 nucleotide mutations and 46 amino acid differences compared to its virulent parental virus, ALD strain (Ishikawa et al., 1995). The GPE- reversion to virulence mapped to three substitutions at positions T830A in E2 glycoprotein and

V2475A/A2563V in NS4B protein.

Residue A830 (present in ALD strain, BICv, and homologous to position 848 in Flag T4v and Flag T4Gv) seems to be critical for CSFV virulence. Interestingly, this residue is one of the five amino acid residues in the T4 mutation of Flag T4v and Flag T4Gv (830AVSPT834 to

830SFNMD834 and 830SFGMD834, respectively). The T830A substitution in E2 acquired during passaging in pigs was shown to reduce viral replication in guinea pig cells while enhancing spreading and replication in pig cells when compared with parental vGPE_ virus. These results suggest that the residue at position 830 in E2 is involved in host tropism and plays a role in cell- to-cell spreading, probably at the level of viral attachment and entry, as it is well known that the glycoproteins E2 and Erns of pestiviruses mediate viral attachment and entry into host cells

(Hulst et al. 1997, Ronecker et al. 2008 and Tscherne et al 2008). Unfortunately, there is only limited information available on the receptors and pathways involved in CSFV binding and

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entry. Moreover, as E2 is exposed on the outer surface of the virus it is well characterized as the most immunodominant protein of CSFV and responsible for inducing neutralizing antibodies in infected pigs (Weiland et al. 1990). As such, amino acid at position 830 is located in the N- terminal region of E2, and this region is the minimal domain required for binding to pig antibodies generated during CSFV infection (Lin et al. 2000 and Zhang et al. 2006).

Consequently, the direct impact of the substitution of alanine for threonine at position 830 in E2 gene on virus infection in swine cells remains unclear and needs to be further examined.

Although mutations observed in NS4B in GPE- revertant were not observed in Flag T4SP revertant virus our lab has also previously demonstrated the importance of this gene in CSFV virulence (Fernandez-Sainz et al., 2010; Gladue et al., 2011). Nevertheless, the amino acid substitutions at positions 2475 and 2563 in NS4B have not yet been reported to be involved in virulence. On the basis of the predicted topology of CSFV NS4B, the residues at positions 2475 and 2563 are located in the N-terminal and C-terminal transmembrane domains, respectively.

The role of transmembrane domains of CSFV NS4B in the virus life cycle is not well understood, while these domains of hepatitis C virus (HCV) are reported to play a role in viral replication (Han et al., 2011). Importantly, we previously identified a putative Toll/interleukin-1 receptor-like domain in the C-terminal region of CSFV NS4B immediately downstream of residue 2563 which resulted in an attenuated phenotype along with enhanced activation of Toll- like receptor 7 (TLR-7)-induced genes (Fernandez-Sainz et al., 2010). We believe this domain is involved in enhanced transcriptional activation of TLR-7-induced genes in swine macrophages, including sustained accumulation of IL-6 mRNA, and is also involved with virus virulence, since amino acid substitutions within this area of the genome lead to complete attenuation of BICv

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(Fernandez-Sainz et al., 2010). Although these residues in NS4B were not affected by back- passing Flag T4Gv in piglets it can all be considered general virulence determinants of CSFV as such, it is apparent that virulence of CSFV is a complex multigenic trait involving different genes and functions evidenced by numerous studies in which mutations of different functional domains in CSFV proteins C, E0, E1, E2, p7 and NS4B reduce pathogenicity in pigs indicating that these genes may be involved in determining CSFV virulence (Fernández-Sainz et al. 2009,

Fernández-Sainz et al. 2010, Fernández-Sainz et al. 2011, Gladue et al. 2010, Gladue et al.

2011a, Gladue et al. 2011b, Gladue et al. 2012, Meyers et al. 1999, Risatti et al. 2007a, Risatti et al. 2007b, Sainz et al. 2008, and Tews et al. 2009) To our knowledge, this report along with the report from Tamura et al., (2012) are the only to have identified specific mutations that result in the reversion of live attenuated vaccine viruses, after successive passages in pigs, back to virulent phenotypes. Remarkably, in both instances the observed reversions partially map to a similar area within the E2 glycoprotein of GPE- and Flag T4v genomes.

Notably, we have shown here that substitutions with synonymous codons in Flag T4Gv provided increased genetic stability of the vaccine virus as five serial passages of Flag T4Gv in naïve piglets did not revert to a virulent phenotype. However, because Flag T4Gv was not detected in tonsil extracts obtained after each of the five successive passages in animals consequently there was no virus available for sequencing. Therefore, it was not possible to determine the presence of any genetic changes in Flag T4Gv throughout the passages in swine.

As a result, it is not clear how the introduced mutations that affect the codon usage in the Flag and T4 regions of the genome increase the genetic stability of this attenuated virus.

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In addition to increased genetic stability, Flag T4Gv is also able to provide protection against lethal CSFV challenge as early as 3 day post vaccination. Although, the ability of the

s innate immune system to interact with CSFV replication has been studied in vitro by׳host several groups (Bensaude et al., 2004, Choi et al., 2004, Dong et al., 2013 and Fernandez-Sainz et al., 2010) the innate response factors mediating early protection remains unknown. To begin to understand the possible mechanisms producing early protection in the absence of an acquired antibody and T cell immune response, the presence and quantification of different components of the innate immune response were assessed for 3 animals vaccinated with Flag T4Gv and euthanized 3 days post immunization, 3 animals infected with WT CSFV and euthanized 3 days post infection and 3 animals vaccinated with Flag T4Gv, challenged 3 days later and euthanized

3 days post challenge using cytokine specific ELISAs and an in-house cytokine RT-qPCR assay.

Interestingly, of all the cytokines tested only, IFN-, showed any significant differences in levels between vaccinated, vaccinated/challenged and WT infected animals. Notably, changes in the expression of this cytokines due to CSFV infection have been reported previously (Magkouras et al., 2008, Fernandez-Sainz et al., 2010). Accordingly, Magkouras et al. (2008) reported CSFV possesses mechanisms that hinder the induction and production of IFNs during CSFV infection.

Furthermore, a direct correlation between the virulence of a CSFV strain and the amount of IFN produced during CSFV infection in swine has also been reported (Renson et al., 2010,

Summerfield et al., 1998 and Tarradas et al., 2014). Likewise, the findings from this study suggest that increased levels of IFN- after Flag T4Gv vaccination may play a protective role against BICv challenge. Definitely, establishment of a robust antiviral state, both systemically and localized to primary sites of virus replication, seems to be crucial to preventing virus replication and spread.

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Recently a live attenuated chimeric BVDV/CSF vaccine, CP7_E2alf has been submitted for licensing for usage in the countries of EU. This vaccine is a BVDV CP7 strain based chimera, expressing CSFV E2 of CSFV Alfort 187. The chimeric virus was characterized in vitro, and the vaccine’s efficacy and marker properties were investigated in immunization/challenge experiments. We believe our Flag T4Gv vaccine will prove to be a better alternative for CSFV control and emergency vaccine usage in countries free of CSFV for two main reasons. First, for CSFV control in the wild boar population of Europe, CP7_E2alf lacks a positive marker which prohibits the possibility to confirm whether analyzed animals have consumed baits, as a result, the effectiveness of CP7_E2alf vaccine campaigns in wild boar cannot be feasibly established. In contrast, our vaccine contains a positive marker, the flag epitope, which makes testing possible by either an ELISA that detects Flag antibodies or by genetic real-time PCR test that recognize the Flag sequence. More importantly, the earliest

CP7_E2alf has been shown to protect animals against lethal CSFV challenge is 7 days post after

IM vaccination, whereas, we have demonstrated that our vaccine protects as early as 3 days post intramuscular vaccination. This is very important for emergency vaccination usage due to the fact that the sooner animals are completely protected from CSFV infection the more likely the virus will be able to be contained to a smaller region thereby limiting potential spreading during an outbreak.

In summary, we have demonstrated that precise mapping and modification of genetic determinants of virulence responsible for reverting a live attenuated strain back to virulence

(Flag T4v), can lead to a more genetically stable attenuated phenotype (Flag T4Gv). More

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importantly, shown here is the development of a CSF DIVA vaccine that could potentially be used for CSFV control in an effort to eradicate CSFV in EU as well as be used for emergency vaccination in CSFV free countries.

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REFERENCES

Agapov, E. V., Murray, C. L., Frolov, I., Qu, L., Myers, T. M., and Rice, C. M. 2004. Uncleaved NS2-3 is required for production of infectious bovine viral diarrhea virus. J. Virol. 78: 2414-25.

Akira, S. and Takeda, K. 2004. Toll-like receptor signalling. Nat. Rev. Immunol. 4: 499-511.

Ahn, J. H., Xu, Y., Jang, W. J., Matunis, M. J., and Hayward, G. S. 2001. Evaluation of interactions of human cytomegalovirus immediate-early IE2 regulatory protein with small ubiquitin-like modifiers and their conjugation enzyme Ubc9. J. Virol. 75: 3859–3872.

Altschul, S. F., Madden, T. L., Schaffer, A. A., Zhang, J., Zhang, Z., Miller, W. and Lipman, D.J. 1997. Gapped BLAST and PSI–BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25: 3389–3402.

Andrew, M. E., Morrissy, C. J., Lenghaus, C., Oke, P. G., Sproat, K. W., Hodgson, A. L. M., Johnson, M. A. and Coupar, B. E. H. 2000. Protection of pigs against classical swine fever with DNA-delivered gp55. Vaccine. 18: 1932–1938.

Ansari, I. H., Kwon, B., Osorio, F. A., and Pattnaik, A. K. 2006. Influence of N-linked glycosylation of porcine reproductive and respiratory syndrome virus GP5 on virus infectivity, antigenicity, and ability to induce neutralizing antibodies. J. Virol. 80: 3994–4004.

Back, N. K., Smit, L., Schutten, M., Nara, P. L., Tersmette, M., and Goudsmit, J. 1993. Mutations in human immunodeficiency virus type 1 gp41 affect sensitivity to neutralization by gp120 antibodies. J. Virol. 67: 6897–6902.

Bauhofer. O., Summerfield, A., Yoshihiro Sakoda, Y., Jon-Duri Tratschin, J., Hofmann, M. A., and Ruggli, N. 2007. Classical Swine Fever Virus Npro Interacts with Interferon Regulatory Factor 3 and Induces Its Proteasomal Degradation. J. Virol. 81: 3087–3096

Becker, P., Avalos Ramirez, R., Orlich, M., Cedillo Rosales, S., Konig, M., Scheizer, H., Stadler, H., Schirmeier, H. and Thiel, H. J. 2003. Genetic and antigenic characterization of novel pestivirus genotypes: implications of classifications. Virol. 311: 96-104.

Beer, M. Reimann, I., Hoffmann, B., and Depner, K. 2007. Novel marker vaccines against classical swine fever. Vaccine. 30: 5665-5670.

Bensaude, E., Turner, J. L. E., Wakeley, P. R., Sweetman, D. A., Pardieu, C., Drew, T. W., Wileman, T. and Powel, P. P. 2004. Classical swine fever virus induces proinflammatory cytokines and tissue factor expression and inhibits apoptosis and interferon synthesis during the establishment of long-term infection of porcine vascular endothelial cells. J. Gen. Virol. 85:1029–1037.

118

Bintintan, I. and Meyers, G. 2010. A New Type of Signal Peptidase Cleavage Site Identified in an RNA Virus Polyprotein. J Biol. Chem. 285: 8572–8584.

Blight, K. J. 2007. Allelic variation in the hepatitis C virus NS4B protein dramatically influences RNA replication. J. Virol. 81: 5724-5736

Bouma, A., de Smit, A. J., de Kluijver, E. P., Terpstra, C. and Moormann R. J. M. 1999. Efficacy and stability of a subunit vaccine based on glycoprotein E2 of classical swine fever virus. Vet. Microbiol. 66: 101-114.

Branza-Nichita, N., Lazar, C., Dwek, R.A., and Zitzmann, N. 2004. Role of N-glycan trimming in the folding and secretion of the pestivirus protein E(rns). Biochem. Biophys. Res. Commun. 319: 655–662.

Brass, V., E. Bieck, R. Montserret, B. Wolk, J. A. Hellings, H. E. Blum, F. Penin, and D. Moradpour. 2002. An amino-terminal amphipathic alpha-helix mediates membrane association of the hepatitis C virus nonstructural protein 5A. J. Biol. Chem. 277: 8130-8139.

Brass, V., Pal, Z., Sapay, N., Deleage, G., Blum, H. E., Penin, F., and Moradpour, D. 2007. Conserved determinants for membrane association of nonstructural protein 5A from hepatitis C virus and related viruses. J. Virol. 81: 2745–2757.

Brock, K. V., Deng, R., and Riblet, S. M. 1992. Nucleotide sequencing of 50 and 30 termini of bovine viral diarrhea virus by RNA ligation and PCR. J. Virol. Methods 38: 39–46.

Carlson, U. and Belak, K. 1994. Border disease virus transmitted to sheep and cattle by a persistently infected ewe: epidemiology and control. Acta Vet. Res. 35: 79-88.

Chang, T. H., Kubota, T., Matsuoka, M., Jones, S., Bradfute, S. B., Bray, M., and Ozato, K. 2009. Ebola Zaire virus blocks type I interferon production by exploiting the host SUMO modification machinery. PLoS Pathog. 5: e1000493.

Chang C. Y., Huang C. C., Lin Y .J., Deng M. C., Chen H. C., Tsai C. H., Chang W. M., and Wang F. I. 2010. Antigenic domains analysis of classical swine fever virus E2 glycoprotein by mutagenesis and conformation-dependent monoclonal antibodies. Virus Res. 149: 183–189.

Chen, Y., Xiao, J., Xiao, J., Sheng, C., Wang, J., Jia, L., Zhi, Y., Li, G., Chen, J., and Xiao, M. 2012. Classical swine fever virus NS5A regulates viral RNA replication through binding to NS5B and 3’UTR. Virol. 432: 376–388.

Chiocca, S. 2007. Viral control of the SUMO pathway: Gam1, a model system. Biochem. Soc. Trans. 35: 1419–1421.

119

Chiu, M. W., Shih, H. M., Yang, T. H., and Yang, Y. L. 2007. The type 2 dengue virus envelope protein interacts with small ubiquitin-like modifier-1 (SUMO-1) conjugating enzyme 9 (Ubc9). J. Biomed. Sci. 14: 429–444.

Choi, C., Whuang, K. K., and Chae, C.. 2004. Clasiscal swine fever induces tumor necorosis factor-α and lymphocyte apoptosis. Arch. Virol., 149: 875–889.

Collett, M. S., Wiskerchen, M. A., Welniak, E., and Belzer, S. K. 1991. Bovine viral diarrhea virus genomic organization. Arch. Virol. Suppl. 3:19–27.

Combet, C., Blanchet, C., Geourjon, C. and Deleage, G. 2000. NPS@: network protein sequence analysis. Trends Biochem. Sci. 25: 147-50.

CORDIS: CSFV_goDIVA Report Summary. c2013. [accessed 2016 July, 1]. http://cordis.europa.eu/result/rcn/53691_en.html.

Dapner K. R., Bouma, A., Koenen, F., Klinkenberg, D., Lange, E., de Smit, H. and Vanderhallen, H. 2001. Classical swine fever (CSF) marker vaccine. Trial II. Challenge study in pregnant sows. Vet. Microbiol. 83: 107–120. de Moerlooze, L., Desport, M., Renard, A., Lecomte, C., Brownlie, J., and Martial, J. A. 1990. The coding region for the 54-kDa protein of several pestiviruses lacks host insertions but reveals a “zincfinger-like” domain. Virol. 177: 812–815. de Smit, A. J., Bouma, A., van Gennip, H. G. P., de Kluijver, E. P. and Moormann, R. J. M. 2001. Chimeric (marker) C-strain viruses induce clinical protection against virulent classical swine fever virus (CSFV) and reduce transmission of CSFV between vaccinated pigs. Vaccine. 19: 1467–1476.

Doms, R. W., Lamb, R. A., Rose, J. K., and Helenius, A. 1993. Folding and assembly of viral membrane proteins. Virol. 193: 545-562

Done, S. H., Potter, R. A., Gresham, A. C. J. and Chennells, D. G. 2000. Post–weaning multisystemic wasting syndrome and porcine dermatitis and nephropathy syndrome – two new pig diseases: a simple view of the clinical indicators. In Practice 23: 14-21.

Dong, X. N., Wei, K., Liu, Z. Q. and Chen, Y. H. 2002. Candidate peptide vaccine induced protection against classical swine fever virus. Vaccine. 21: 167–173.

Dong, X. Y., Liu, W. J., Zhao, M. Q., Wang, J. Y., Pei, J. J., Luo, Y. W., Ju, C. M., and Chen, J. D.. 2013. Classical swine fever virus triggers RIG-I and MDA5-dependent signaling pathway to IRF-3 and NF-κB activation to promote secretion of interferon and inflammatory cytokines in porcine alveolar macrophages. Virol. J., 10: 286.

120

Donis, R. O. and Dubovi, E. J. 1987. Molecular specificity of the antibody responses of cattle naturally and experimentally infected with cytopathic and noncytopathic bovine viral diarrhea virus biotypes. Am. J. Vet. Res. 48: 1549–1554.

Edwards, S., Moennig, V. and Wensvoort, G. 1991. The development of an international reference panel of monoclonal antibodies for the differentiation of hog cholera virus from other pestiviruses. Vet. Microbiol. 29: 101-108.

Edwards, S., Fukusho, A., Lefevre, P., Lipowski, A., Pejsak, Z., Roeche, P. and Westergaard, J. 2000. Classical swine fever: the global situation. Vet. Microbiol. 73: 103- 119.

Elbers, K.,Tautz, N., Becher,P., Rumenapf, T., and Thiel, H. J. 1996. Processing in the Pestivirus E2-NS2 region: Identification of the non-structural proteins p7and E2p7. Journal of Virol. 70: 4131–4135.

El Omari, K., Iourin, O., Harlos, K., Grimes, J. M., and Stuart, D. I. 2013. Structure of a Pestivirus Envelope Glycoprotein E2 Clarifies Its Role in Cell Entry. Cell Rep. 3: 30–35.

Elazar, M., Liu, P., Rice, C. M. and Glenn J. S. 2004. An N-terminal amphipathic helix in hepatitis C virus (HCV) NS4B mediates membrane association, correct localization of replication complex proteins, and HCV RNA replication. J. Virol. 78:11393-11400

Einav, S., Elazar, M., Danieli, T., and Glenn, J. S. 2004. A nucleotide binding motif in hepatitis C virus (HCV) NS4B mediates HCV RNA replication. J. Virol. 78: 11288–11295.

Fernandez Sainz, I., Holinka, L. G., Lu, Z., Risatti, G. R., and Borca, M. V. 2008. Removal of a N-linked glycosylation site of classical swine fever virus strain Brescia Erns glycoprotein affects virulence in swine. Virol. 370: 122-129

Fernandez-Sainz, I., Holinka, L. G., Gavrilov, B. K., Prarat, M. V., Gladue, D., Lu, Z., Jia, W., Risatti, G. R., and Borca, M. V. 2009. Alteration of the N-linked glycosylation condition in E1 glycoprotein of Classical Swine Fever Virus strain Brescia alters virulence in swine. Virol. 386: 210–216.

Fernandez-Sainz, I., Gladue, D. P., Holinka, L. G., O'Donnell, V., Gudmundsdottir, I., Prarat, M. V., Patch, J. R., Golde, W. T., Lu, Z., Zhu, J., Carrillo, C., Risatti, G. R., and Borca, M. V. 2010. Mutations in classical swine fever virus NS4B affect virulence in swine. J. Virol. 84: 1536–1549

Fernandez-Sainz, I. J., Holinka, L. G., Gladue, D. P., O’Donell, V., Lu, Z., Gavrilov, B. K., Risatti, G. R., and Borca, M. V. 2011. Substitution of specific cysteine residues in the E1 glycoprotein of classical swine fever virus strain Brescia affects formation of E1-E2 heterodimers and alters virulence in swine. J. Virol. 85: 7264-7272

121

Fernández-Sainz, I. J., Largo, E., Gladue, D. P., Fletcher, P., O’Donnell, V.O., Holinka, L. G., Carey, L. B., Lu, Z., Nieva, J. L., and Borca, M. V. 2014 Effect of specific amino acid substitutions in the putative fusion peptide of structural glycoprotein E2 on Classical Swine Fever Virus replication. Virol 457: 121-130

Fetzer, C., Tews, B. A., and Meyers, G. 2005. The carboxy-terminal sequence of the pestivirus glycoprotein E(rns) represents an unusual type of membrane anchor. J. Virol. 79: 11901–11913.

Fiebach, A.R., Guzylack-Piriou, L., Python, S., Summerfield, A., and Ruggli, N. 2011. Classical Swine Fever virus npro limits type I interferon induction in plasmacytoid dendritic cells by interacting with interferon regulatory factor 7. J Virol. 85: 8002–8011.

Fournillier, A., Wychowski, C., Boucreux, D., Baumert, T. F., Meunier, J. C., Jacobs, D., Muguet, S., Depla, E., and Inchauspe, G. 2001. Induction of hepatitis C virus E1 envelope protein-specific immune response can be enhanced by mutation of Nglycosylation sites. J. Virol. 75: 12088–12097.

Fukata, M., Watanabe, T., Noritake, J., Nakagawa, M., Yamaga, M., Kuroda, S., Matsuura, Y., Iwamatsu, A., Perez, F., and Kaibuchi, K. 2002. Rac1 and Cdc42 capture microtubules through IQGAP1 and CLIP-170. Cell 109: 873–885.

Garry, R. F., and Dash, S. 2003. Proteomics computational analyses suggest that hepatitis C virus E1 and pestivirus E2 envelope glycoproteins are truncated class II fusion proteins. Virol. 307: 255–265.

Gavel, Y., and G. von Heijne. 1990. Sequence differences between glycosylated and nonglycosylated Asn-X-Thr/Ser acceptor sites: implications for protein engineering. Protein Eng. 3: 433–442.

Gavrilov, B. K., Rogers, K., Fernandez-Sainz, I. J., Holinka, L. G. Borca, M. V., and Risatti, G. R. 2011. Effects of glycosylation on antigenicity and immunogenicity of classical swine fever virus envelope proteins. Virol. 420: 135-145

Genovesi, E. V., Villinger, F., Gerstner, D. J., Whyard, T. C. and Knudsen, R. C. 1990. Effect of macrophage-specific colony-stimulating factor (CSF-1) on swine monocyte/macrophage susceptibility to in vitro infection by African swine fever virus. Vet Microbiol. 25: 153–176.

Gladue, D., Holinka, L. G., Fernandez-Sainz, I. J., Prarat, M. V. Lu, Z., Risatti, G. R., and Borca, M. V. 2010. Effects of the interactions of classical swine fever virus Core protein with proteins of the SUMOylation pathway on virulence in swine. Virol. 407: 129-136.

122

Gladue, D., Holinka, L. G., Fernandez-Sainz, I. J., Prarat, M. V. O’Donell, V., Lu, Z., Vepkhvadze, N., Risatti, G. R., and Borca, M. V. 2011. Interaction between Core protein of classical swine fever virus with cellular IQGAP1 protein appears essential for virulence in swine. Virol. 412: 68-74.

Gladue, D.P., Gavrilov, B. K., Holinka, L. G., Fernandez-Sainz, I. J.,Vepkhvadze, N. G., Rogers, K., Risatti, G., and Borca, M. V. 2011. Identification of an NTPase motif in classical swine fever virus NS4B protein. Virol. 411: 41–49.

Gladue, D. P., Holinka,L. G., Largo, E., Fernandez-Sainz, I., Carrillo, C., O’Donnell, V., and Borca, M. V. 2012. Classical swine fever virus p7 protein is a viroporin involved in virulence in swine. J. Virol. 86: 6778–6791.

Gladue, D., O’Donell, V., Fernandez-Sainz, I. J., Fletcher, P., Baker-Branstetter, R., Holinka, L. G., Sanford, B., Carlson, J. Lu, Z., and Borca, M. V. 2014a. Interaction of structural Core protein of Classical Swine Fever Virus with endoplasmic reticulum-associated degradation pathway protein OS9. Virol. 461: 173-179.

Gladue, D., Baker-Branstetter, R., Holinka, L. G., Fernandez-Sainz, I. J., O’Donell, V., Fletcher, P., Lu, Z., and Borca, M. V. 2014b. Interaction of CSFV E2 Protein with Swine Host Factors as Detected by Yeast Two-Hybrid System. PLoS One 9: e85324

Gómez-Villamandos, J. C., Ruiz-Villamor, E., Bautista, M. J., Sánchez, C. P., Sánchez- Cordón, P. J., Salguero, F. J. and Jover, A. 2001. Morphological and immunohistochemical changes in splenic macrophages of pigs infected with classical swine fever. J. Comp. Path. 125: 98–109.

Gong, Y., Trowbridge, R., Macnaughton, T. B., Westaway, E. G., Shannon, A. D., and Gowans, E. J. 1996. Characterization of RNA synthesis during a one-step growth curve and of the replication mechanism of bovine viral diarrhea virus. J. Gen. Virol. 77: 2729–2736

Gosert,R., Egger, D., Lohmann, V., Bartenschlager, R., Blum, H. E., Bienz, K., and Moradpour, D. 2003. Identification of the hepatitis C virus RNA replication complex in huh-7 cells harboring subgenomic replicons. J. Virol. 77: 5487–5492

Gottipati, K., Ruggli, N., Gerber, M., Tratschin, J., Benning, M., Bellamy, H., and Choi, K. H. 2013. The structure of classical swine fever virus N(pro): A novel cysteine Auto-protease and zinc-binding protein involved in subversion of type I interferon induction. PLoS Pathogens 9(10), e1003704: 1 – 10.

Gouttenoire, J., V. Castet, R. Montserret, N. Arora, V. Raussens, J. M. Ruysschaert, E. Diesis, H. E. Blum, F. Penin, and D. Moradpour. 2009. Identification of a novel determinant for membrane association in hepatitis C virus nonstructural protein 4B. J. Virol. 83:6257-6268.

123

Grassmann, C. W., Isken, O., Tautz, N., and Behrens, S. E. 2001. Genetic analysis of the pestivirus nonstructural coding region: Defects in the NS5A unit can be complemented in trans. J. Viro. 75: 7791–7802.

Grassmann, C. W., Yu, H., Isken, O., and Behrens, S. E. 2005. Hepatitis C virus and the related bovine viral diarrhea virus considerably differ in the functional organization of the 50 non-translated region: Implications for the viral life cycle. Virol. 333: 349–366.

Griffin, S. D., Beales, L. P., Clarke, D. S., Worsfold, O., Evans, S. D., Jaeger, J., Harris M. P., and Rowlands D. J. 2003. The p7 protein of hepatitis C virus forms an ion channel that is blocked by the antiviral drug, Amantadine. FEBS Lett. 535: 34–38.

Gu, B., Liu, C., Lin-Goerke, J., Maley, D. R., Gutshall, L. L., Feltenberger, C. A., and Del Vecchio, A. M. 2000. The RNA helicase and nucleotide triphosphatase activities of the bovine viral diarrhea virus NS3 protein are essential for viral replication. J. Virol. 74: 1794–1800.

Hammond, J. M., McCoy, R. J., Jansen, E. S., Morrissy, C. J., Hodgson, A. L. and Johnson, M. A. 2000. Vaccination with a single dose of a recombinant porcine adenovirus expressing the classical swine fever virus gp55 (E2) gene protects pigs against classical swine fever. Vaccine 18: 1040-1050.

Han Q., Aligo J., Manna D., Belton K., Chintapalli S. V., Hong Y., Patterson R. L., van Rossum D. B., and Konan K. V. 2011. Conserved GXXXG- and S/T-like motifs in the transmembrane domains of NS4B protein are required for hepatitis C virus replication. J. Virol. 85: 6464-6479.

Harada, T., Tautz, N. and Thiel, H. 2000. E2-p7 region of the bovine viral diarrhea virus polyprotein: processing and functional studies. J. Virol. 74: 9494-9506.

Heimann, M., G. Roman-Sosa, B. Martoglio, H. J. Thiel, and T. Rumenapf. 2006. Core protein of pestiviruses is processed at the C terminus by signal peptide peptidase. J. Virol. 80:1915-1921

Helenius, A., and M. Aebi. 2001. Intracellular functions of N-linked glycans. Science 291:2364-2369.

Helle, F., Vieyres, G., Elkrief, L., Popescu, C.I., Wychowski, C., Descamps, V., Castelain, S., Roingeard, P., Duverlie, G., and Dubuisson, J. 2010. Role of N-linked glycans in the functions of hepatitis C virus envelope proteins incorporated into infectious virions. J. Virol. 84: 11905–11915.

Hinnebusch, A. G., and Lorsch, J. R. 2012. The mechanism of eukaryotic translation initiation: New insights and challenges. Cold Spring Harb Perspect Biol. 4:10.

124

Holinka L. G., Fernandez-Sainz I., Sanford B., O'Donnell V., Gladue D. P., Carlson J., Lu Z., Risatti G. R., and Borca, M. V. 2014. Development of an improved live attenuated antigenic marker CSF vaccine strain candidate with an increased genetic stability. Virol. 471- 473: 13-18

Holliger P. and Riechmann, L. 1997. A conserved infection pathway for filamentous bacteriophages is suggested by the structure of the membrane penetration domain of the minor coat protein g3p from phage fd. Structure 15: 265-75.

Huh, Y. H., Kim, Y. E., Kim, E. T., Park, J. J., Song, M. J., Zhu, H., Hayward, G. S., and Ahn, J. H. 2008. Binding STAT2 by the acidic domain of human cytomegalovirus IE1 promotes viral growth and is negatively regulated by SUMO. J. Virol. 82: 10444–10454.

Hulst, M. M., Westra, D. F., Wensvoort, G. and Moormann, R. J. M. 1993. Glycoprotein E1 of hog cholera virus expressed in insect cells protects swine from hog cholera. J. Virol. 67: 5435–5442.

Hulst, M. M. and Moormann R. J. 1997. Inhibition of pestivirus infection in cell culture by envelope proteins E(rns) and E2 of classical swine fever virus: E(rns) and E2 interact with different receptors. J. Gen. Virol. 78: 2779–2787.

Hulst, M. M., van Gennip, H. G., and Moormann, R. J. 2000. Passage of classical swine fever virus in cultured swine kidney cells selects virus variants that bind to heparin sulfate due to a single amino acid change in envelope proteinE(rns). J. Virol. 74: 9553–9561.

Ishido, S., Fujita, T., and Hotta, H. 1998. Complex Formation of NS5B with NS3 and NS4A Proteins of Hepatitis C Virus. Biochem. Biophys. Res. Commun. 244: 35–40

Isken, O., Langerwisch, U., Schonherr, R., Lamp, B., Schroder, K., Duden, R., Rümenapf, T. H., and Tautz, N. 2014. Functional characterization of bovine viral diarrhea virus nonstructural protein 5A by reverse genetic analysis and live cell imaging. J. Virol. 88: 82–98.

Ivanyi-Nagy, R., Lavergne, J. P., Gabus, C., Ficheux, D., and Darlix, J. L. 2008. RNA chaperoning and intrinsic disorder in the core proteins of Flaviviridae. Nucleic Acids Research 36: 712–725.

Jablonski, S. A., Luo, M. and Morrow, C. D. 1991. Enzymatic activity of poliovirus RNA polymerase mutants with single amino acid changes in the conserved YGDD amino acid motif. J. Virol. 65: 4565–4572.

Kaplan, C. 1989. Vaccinia virus: a suitable vehicle for recombinant vaccines? Arch. Virol. 106: 127-139.

Kerscher, O. 2007. SUMO junction—what's your function? New insights through SUMO-interacting motifs. EMBO Rep. 8: 550–555.

125

Koenig, P., Lange, E., Reimann, I., and Beer M. 2007. CP7_E2alf: a safe and efficient marker vaccine strain for oral immunisation of wild boar against Classical swine fever virus (CSFV). Vaccine. 17: 3391-3339.

Konig, M., Lengsfeld, T., Pauly, T., Stark, R. and Thiel H. J. 1995. Classical swine fever virus: independent induction of protective immunity by two structural glycoproteins. J. Virol. 69: 6479–6486.

Krey, T., Bontems, F.,Vonrhein, C., Vaney, M. C., Bricogne, G., Rumenapf, T., and Rey, F. A. 2012. Crystal structure of the pestivirus envelope glycoprotein E(rns) and mechanistic analysis of its ribonuclease activity. Structure 20: 862–873.

Kummerer, B. M. and Meyers, G. 2000. Correlation between point mutations in NS2 and the viability and cytopathogenicity of bovine viral diarrhea virus strain Oregon analyzed with an infectious cDNA clone. J. Virol. 74: 390–400.

Lackner, T., Muller, A., Pankraz, A., Becher, P., Thiel, H. J., Gorbalenya, A. E., et al. 2004. Temporal modulation of an autoprotease is crucial for replication and pathogenicity of an RNA virus. J. Virol. 78: 10765–10775.

Lackner, T., Thiel, H. J., and Tautz, N. 2006. Dissection of a viral autoprotease elucidates a function of a cellular chaperone in proteolysis. Proceedings of the National academy of Sciences of the United States of America. 103: 1510–1515.

Langedijk, J. P., van Veelen, P. A. Schaaper, W. M. de Ru, A. H. Meloen, R. H. and Hulst, M. M. 2002. A structural model of pestivirus E(rns) based on disulfide bond connectivity and homology modeling reveals an extremely rare vicinal disulfide. J. Virol. 76: 10383-10392

Largo, E., Gladue, D. P., Huarte, N., Borca, M. V., and Nieva, J. L. 2014. Pore-forming activity of pestivirus p7 in a minimal model system supports genus-specific viroporin function. Antiviral Res. 101: 30–36.

Lee, J., T. H. Chuang, V. Redecke, L. She, P. M. Pitha, D. A. Carson, E. Raz, and H. B. Cottam. 2003. Molecular basis for the immunostimulatory activity of guanine nucleoside analogs: activation of Toll-like receptor 7. Proc. Natl. Acad. Sci. U. S. A. 100:6646-6651.

Leifer I., Lange E., Reimann I., Blome S., Juanola S., Duran J. P., and Beer M. 2009. Modified live marker vaccine candidate CP7_E2alf provides early onset of protection against lethal challenge infection with classical swine fever virus after both intramuscular and oral immunization. Vaccine. 27: 6522-6529.

Liess, B. 1987. Pathogenesis and epidemiology of hog cholera. Ann. Rech. Vet. 18: 139-145.

Letunic I., Copley R. R., Pils B., Pinkert S., Schultz J., and Bork P. 2006. SMART 5: domains in the context of genomes and networks. Nucleic Acids Res. 34 (Database issue):D257- 260

126

Leung, J., Yueh, A., Appah Jr., F. S., Yuan, B., de los Santos, K., and Goff, S. P. 2006. Interaction of Moloney murine leukemia virus matrix protein with IQGAP. EMBO J. 25: 2155– 2166.

Leyssen, P., Clercq, E., and Neyts, J. 2000. Perspectives for the treatment of infections with flaviviridae. Clin. Microbiol. Rev. 13: 67-82.

Li, Y., Wang, J., Kanai, R., and Modis, Y. 2013. Crystal structure of glycoprotein E2 from bovine viral diarrhea virus. Proceedings of the National Academy of Sciences of the United States of America. 110: 6805–6810.

Li, L, Wu, R, Zheng, F., Zhao, C., Pan, Z. 2015. The N-terminus of classical swine fever virus (CSFV) nonstructural protein 2 modulates viral genome RNA replication. Virus Res. 210: 90-99.

Liang, D., Fernandez Sainz, I., Ansari, I. H., Gil, L. H. V. G., Vassilev, V. And Donis, R. 2003. The envelope glycoprotein E2 is a determinant of cell culture tropism in ruminant pestiviruses. J. Gen. Virol. 84: 1269-1274.

Liang, D., Chen, L., Ansari, I. H., Gil, L. H., Topliff, C. L., Kelling, C. L., and Donis R. O. 2009. A replicon trans-packaging system reveals the requirement of nonstructural proteins for the assembly of bovine viral diarrhea virus (BVDV) virion. Virol. 387: 331–340.

Lin, C., Wu, J. W., Hsiao, K., and Su, M. S. 1997. The hepatitis C virus NS4A protein: interactions with the NS4B and NS5A proteins. J. Virol. 71: 6465–6471.

Lin, M., Lin, F., Mallory, M., and Clavijo, A. 2000. Deletions of structural glycoprotein E2 of classical swine fever virus strain alfort/187 resolve a linear epitope of monoclonal antibody WH303 and the minimal N-terminal domain essential for binding immunoglobulin G antibodies of a pig hyperimmune serum. J. Virol. 74: 11619–11625.

Lindenbach, B. D. and Rice, C. M. 2001. Flaviviridae: the viruses and their replication, p.991- 1042. In B.N. Fields, D.M. Knipe, and P. M. Howley (ed), Fields Virology, 4th ed. Lippincott Williams & Wilkins, Philadelphia, Pa.

Luo, X., Ling, D., Li, T., Wan, C., Zhang, C., Pan, Z. 2009. Classical swine fever virus Erns glycoprotein antagonizes induction of interferon-by double-stranded RNA. Can. J. Microbiol. 55: 698–704.

Lundin, M., Monne, M., Widell, A., Von Heijne, G. and Persson, M. A. 2003. Topology of the membrane-associated hepatitis C virus protein NS4B. J. Virol. 77: 5428-5438

Ma, Y., Yates, J., Liang, Y., Lemon, S. M., and Yi, M. 2008. NS3 helicase domains involved in infectious intracellular hepatitis C virus particle assembly. J. Virol. 82: 7624–7639.

127

Magkouras, I., Mätzener, P., Rümenapf, T., Peterhans, E., and Schweizer, M. 2008. RNase- dependent inhibition of extracellular, but not intracellular, dsRNA-induced interferon synthesis by Erns of pestiviruses. J. Gen. Virol., 89: 2501–2506.

Maurer, R., Stettler, P., Ruggli, N., Martin A. Hofmann, M. A. and Tratschin, J. 2005. Oronasal vaccination with classical swine fever virus (CSFV) replicon particles with either partial or complete deletion of the E2 gene induces partial protection against lethal challenge with highly virulent CSFV. Vaccine. 23: 3318–3328

Mayer, D., Hofmann M. A. and Tratschin, J. D. 2004. Attenuation of classical swine fever virus by deletion of the viral N(pro) gene. Vaccine 22: 317-328.

Meyers, G. and Thiel, H. J. 1996. Molecular characterization of pestiviruses. Adv. Vir. Res. 47: 53-118.

Meyers, G., Saalmuller, A. and Buttner M. 1999. Mutations abrogating the RNase activity in glycoprotein E(rns) of the pestivirus classical swine fever virus lead to virus attenuation. J. Virol. 73: 10224-10235.

Migration Policy Institute: Frequently Requested Statistics on Immigrants and Immigration in the United States. c2014. [accessed 2016 June 07]. http://www.migrationpolicy.org/article/ frequently-requested-statistics-immigrants-and-immigration-united-states.

Mittelholzer, C., Moser, C., Tratschin, J. D. and Hofmann, M. A. 2000. Analysis of classical swine fever virus replication kinetics allows differentiation of highly virulent from avirulent strains. Vet. Microbiolog. 74: 5463-5467

Moennig, V., Floegel-Niesmann, G. and Greiser-Wile, I. 2003. Clinical signs and epidemiology of classical swine fever: a review of new knowledge. Vet. Journal. 165: 11-20.

Moormann, R. J., Warmerdam, P.A., van der Meer, B., and Hulst, M. M. 1990. Nucleotide sequence of hog cholera virus RNA: properties of the polyprotein encoded by the open reading frame spanning the viral genomic RNA. Vet. Microbiol. 23:185–191.

Moormann, R. J. M., Bouma A., Kramps, J. A., Terpstra, C. and de Smit, H. J. 2000. Development of classical swine fever subunit marker vaccine and companion diagnostic test. Vet. Microbiol. 73: 209 – 219.

Moes, L., and Wirth, M. 2007. The internal initiation of translation in bovine viral diarrhea virus RNA depends on the presence of an RNA pseudoknot upstream of the initiation codon. Virol. J. 4: 124.

Moulin, H. R., Seuberlich, T., Bauhofer, O., Bennett, L.C., Tratschin, J. D., Hofmann, M. A., et al. 2007. Nonstructural proteins NS2-3 and NS4A of classical swine fever virus: Essential features for infectious particle formation. Virol. 365: 376–389.

128

Mukhopadhyay, S., Kuhn, R. J., and Rossmann, M. G. 2005. A structural perspective of the flavivirus life cycle. Nature Reviews 3: 13-22.

Mulder, N. J., Apweiler, R., Attwood, T. K., Bairoch, A., Bateman, A., Binns, D., Bradley, P., Bork, P., Bucher, P., Cerutti, L., Copley, R., Courcelle, E., Das, U., Durbin, R., Fleischmann, W., Gough, J., Haft, D., Harte, N., Hulo, N., Kahn, D., Kanapin, A., Krestyaninova, M., Lonsdale, D., Lopez, R., Letunic, I., Madera, M., Maslen, J., McDowall, J., Mitchell, A., Nikolskaya, A. N., Orchard, S., Pagni, M., Ponting, C. P., Quevillon, E., Seeler, J. S., and Dejean, A. 2003. Nuclear and unclear functions of SUMO. Nat. Rev. Mol. Cell Biol. 4: 690–699.

Muñoz-Jordan, J. L., Sanchez-Burgos, G. G. Laurent-Rolle, M., and Garcia-Sastre A. 2003. Inhibition of interferon signaling by dengue virus. Proc. Natl. Acad. Sci. U. S. A. 100: 14333-14338.

Murray, C. L., Marcotrigiano, J., and Rice, C. M. 2008. Bovine viral diarrhea virus core is an intrinsically disordered protein that binds RNA. J. Virol. 82: 1294–1304.

Niedenthal, R. 2007. UBC-9 fusion-directed SUMOylation (UFDS). Biochem. Soc. Trans. 35: 1430–1432. Office of Travel & Tourism: I-94 Program: 2014 Monthly Arrivals Data. c2014. [accessed 2016 June 07]. http://travel.trade.gov/view/m-2014-I-001/index.html.

OIE World Organisation for Animal Health: OIE listed diseases 2016. c2016. [accessed 2016 April 25]. http://www.oie.int/en/animal-health-in-the-world/oie-listed-diseases-2016.

O'Reilly E. K. and Kao, C. C. 1998. Analysis of RNA-dependent RNA polymerase structure and function as guided by known polymerase structures and computer predictions of secondary structure. Virol. 252: 287 -303.

Palacios, S., Perez, L. H., Welsch, S., Schleich, S., Chmielarska, K., Melchior, F., and Locker, J. K. 2005. Quantitative SUMO-1 modification of a vaccinia virus protein is required for its specific localization and prevents its self-association. Mol. Biol. Cell 16: 2822–2835.

Patkar, C. G.,and Kuhn,R. J. 2008. Yellow Fever virus NS3 plays an essential role in virus assembly independent of its known enzymatic functions. J. Virol. 82: 3342–3352.

Paton, D. J. and Greiser-Wilke, I. 2003. Classical swine fever – an update. Res. Vet. Sci. 75: 169-178.

Pauly, T., Konig, M., Thiel, H. J. and Saalmuller, A. 1998. Infection with classical swine fever virus: effects on phenotype and immune responsiveness of porcine T lymphocytes. J. Gen. Virol. 79: 31–40.

129

Piccininni, S., Varaklioti, A., Nardelli, M., Dave, B., Raney, K. D., McCarthy, J. E. 2002. Modulation of the hepatitis C virus RNA dependent RNA polymerase activity by the non structural (NS) 3 helicase and the NS4B membrane protein. J. Biol. Chem. 277:45670–45679.

Poole, T. L., Wang, C., Popp, R. A., Potgieter, L. N. D., Siddiqui, A., and Collett, M. S. 1995. Pestivirus translation occurs by internal ribosome entry. Virology 206: 750–754.

Qu, L., McMullan, L. K., and Rice C. M. 2001. Isolation and characterization of noncytopathic pestivirus mutants reveals a role for nonstructural protein NS4B in viral cytopathogenicity. J. Virol. 75: 10651-10662.

Rawlings, N. D., Barrett, A. J., and Bateman, A. 2012. MEROPS: The database of proteolytic enzymes, their substrates and inhibitors. Nucleic Acids Res. 40: (Database issue), D343– D350.

Reed, L. J. and Muench, H. A. 1938. A simple method of estimating fifty percent endpoints. Am. J. Hyg. 27: 493-497.

Reed, K. E., Gorbalenya, A. E., and Rice, C .M. 1998. The NS5A/NS5 proteins of viruses from three genera of the family flaviviridae are phosphorylated by associated serine/ threonine kinases. J. Virol. 72: 6199–6206.

Reimann I., Depner K., Trapp S., and Beer M. 2004. An avirulent chimeric Pestivirus with altered cell tropism protects pigs against lethal infection with classical swine fever virus. Virol. 322: 143–157.

Reitter, J. N., Means, R. E., and Desrosiers, R. C. 1998. A role for carbohydrates in immune evasion in AIDS. Nat. Med. 4 : 679–684.

Renson, P., Blanchard, Y., Le Dimna, M., Felix, H., Cariolet, R., Jestin, A., Le Potier, M. F.. 2010. Acute induction of cell death-related IFN stimulated genes (ISG) differentiates highly from moderately virulent CSFV strains. Vet. Res. 41: 7.

Renson, P., Le Dimna, M., Keranflech, A., Cariolet, R., Koenen, F., and Le Potier, M. F. 2013. CP7_E2alf oral vaccination confers partial protection against early classical swine fever virus challenge and interferes with pathogeny-related cytokine responses. Vet. Res. 44: 9.

Rice, C. M. (1996). Flaviviridae: the viruses and their replication. In “Fields Virology” (B.N. Fields, D.M. Knipe, and P. M. Howley, Eds.) 3rd Ed., Lippincott-Raven, Philadelphia, Pa.

Riedel, C., Lamp, B., Heimann, M., and Rumenapf, T. 2010. Characterization of Essential Domains and Plasticity of the Classical Swine Fever Virus Core Protein. J. Virol. 84: 11523– 11531.

130

Riedel, C., Lamp, B., Heimann, M., Konig, M., Blome, S., Moennig, V., Schuttler, C., Thiel, H., and Rumenapf, T. 2012. The Core Protein of Classical Swine Fever Virus Is Dispensable for Virus Propagation In Vitro. PloS Pathog. 8: e1002598

Rinck, G., Birghan, C., Harada, T., Meyers, G., Thiel, H.J., and Tautz, N. 2001. A cellular J-domain protein modulates polyprotein processing and cytopathogenicity of a pestivirus. J. Virol. 75: 9470-9482.

Risatti, G. R., Callahan, J. D., Nelson, W. M. and Borca, M. V. 2003. Rapid detection of classical swine fever virus by a portable real-time reverse transcriptase PCR assay. J. Clin. Microbiol. 41: 500-505

Risatti, G. R., Borca, M. V., Kutish, G. F., Lu, Z., Holinka, L. G., French R. A., Tulman E. R. and Rock D. L. 2005a. The E2 glycoprotein of classical swine fever virus is a virulence determinant in swine. J. Virol. 79: 3787-3796.

Risatti, G. R., Holinka, L. G., Lu, Z., Kutish, G. F., Tulman, E. R., French, R. A., Sur, J. H., Rock, D. L., and Borca, M. V. 2005b. Mutation of E1 glycoprotein of classical swine fever affects viral virulence in swine. Virol. 343: 116-127.

Risatti, G. R., Holinka, L. G., Carrillo, C., Kutish, G. F., Lu, Z., Tulman, E. R., Sainz, I. F. and Borca, M. V. 2006. Identification of a novel virulence determinant within the E2 structural glycoprotein of classical swine fever virus. Virol. 355: 94-101.

Risatti, G. R., Holinka, L. G., Fernandez Sainz, I., Carrillo, C., Lu, .Z. and Borca, M. V. 2007. N-linked glycosylation status of Classical Swine Fever Virus strain Brescia E2 glycoprotein influences virulence in swine. J. Virol. 81: 924-933

Rumenapf, T., Stark, R., Meyers, G. and Thiel H. J. 1991. Structural proteins of hog cholera virus expressed by vaccinia virus: further characterization and induction of protective immunity. J. Virol. 65: 589–597.

Rumenapf, T. Unger, G., Strauss, J. H. and Thiel, H. J. 1993. Processing of the envelope glycoproteins of pestiviruses. J. Virol. 67: 3288-3294.

Rumenapf, T., Stark, R., Heimann, M., and Thiel, H. J. 1998. N-terminal protease of pestiviruses: Identification of putative catalytic residues by site-directed mutagenesis. J. Virol. 72: 2544–2547.

Sadanari, H., Yamada, R., Ohnishi, K., Matsubara, K., and Tanaka, J. 2005. SUMO-1 modification of the major immediate-early (IE) 1 and 2 proteins of human cytomegalovirus is regulated by different mechanisms and modulates the intracellular localization of the IE1, but not IE2, protein. Arch. Virol. 150: 1763–1782

131

Sanger, F., Nickelson, S. and Coulson A. R. 1977. DNA sequencing with chain-terminating inhibitors. Proc, Natl. Acad. Sci. USA 74: 5463-5467.Sapay N, Montserret R, Chipot C, Brass V, Moradpour D, Deleage G, Penin F. 2006. NMR structure and molecular dynamics of the in-plane membrane anchor of nonstructural protein 5A from bovine viral diarrhea virus. Biochem. 45: 2221–2233.

Schneider, R., Unger, G., Stark, R., Schneider-Scherzer, E., and Thiel, H. J. 1993. Identification of a structural glycoprotein of an RNA virus as a ribonuclease. Science 261: 1169-1171.

Scott C., and Griffin S.D. 2015. Viroporins: Structure, function and potential as antiviral targets. J. Gen. Virol. 96: 2000-27.

Seeler, J. S., and Dejean, A. 2003. Nuclear and unclear functions of SUMO. Nat. Rev. Mol. Cell Biol. 4: 690–699.

Selengut, J., Sigrist, C. J., Silventoinen, V., Studholme, D. J., Vaughan, R. and Wu, C. H. 2005. InterPro, progress and status in 2005. Nucleic Acids Res. 33(Database Issue):D201-205.

Shakin-Eshleman, S. H., A. T. Remaley, J. R. Eshleman, W. H. Wunner, and S. L. Spitalnik. 1992. N-linked glycosylation of rabies virus glycoprotein. Individual sequons differ in their glycosylation efficiencies and influence on cell surface expression. J. Biol. Chem. 267: 10690-10698.

Sheng C., Zhu Z. L., Yu J. L., Wan L. Z., Wang Y. J., Chen J., Gu F. K., and Xiao M. 2010. Characterization of NS3, NS5A and NS5B of classical swine fever virus through mutation and complementation analysis. Vet. Microbiol. 140: 72–80.

Sheng, C., Chen, Y., Xiao, J., Xiao, J., Wang, J., Li, G., Chen, J., and Xiao, M. 2012. Classical swine fever virus NS5A protein interacts with 3’-untranslated region and regulates viral RNA synthesis. Virus Res. 163: 636–643.

Shi, X., and Elliot, R. M. 2004. Analysis of N-linked glycosylation of hantaan virus glycoproteins and the role of oligosaccharide side chains in protein folding and intracellular trafficking. J. Virol. 78: 5414-5422.

Shirai, C., and Mizuta, K. 2008. SUMO mediates interaction of Ebp2p, the yeast homolog of Epstein–Barr virus nuclear antigen 1-binding protein 2, with a RING finger protein Ris1p. Biosci. Biotechnol. Biochem. 72: 1881–1886.

Shirota, Y., Luo, H., Qin, W., Kaneko, S., Yamashita, T., Kobayashi, K., and Murakami, S. 2002. Hepatitis C virus (HCV) NS5A binds RNA-dependent RNA polymerase (RdRP) NS5B and modulates RNA-dependent RNA polymerase activity. J. Biol. Chem. 277: 11149–11155.

132

Stark, R., Meyers, G., Rumenapf, T. and Thiel, H. J. 1993. Processing of pestivirus polyprotein: cleavage site between autoprotease and nucleocapsid protein of classical swine fever virus. J. Virol. 67: 7088-7095.

Steffens, S., Thiel, H. J., and Behrens, S. E. 1999. The RNA-dependent RNA polymerases of different members of the family Flaviviridae exhibit similar properties in vitro. J. Gen. Virol. 80: 2583–2590.

Stegeman, A., Elbers, A., de Smit, H., Moser, H., Smak, J. and Pluimers, F. 2000. The 1997 – 1998 epidemic of classical swine fever in the Netherlands. Vet. Microbiol. 73: 183-196.

Steinmann E, Penin, F., Kallis, S., Patel, A. H., Bartenschlager, R., and Pietschmann, T. 2007. Hepatitis C virus p7 protein is crucial for assembly and release of infectious virions. PLoS Pathog. 3: e103.

Summerfield, A., Knötig, S. M., and McCullough, K. C. 1998. Lymphocyte apoptosis during classical swine fever: implication of activation-induced cell death. J. Virol. 72: 1853–1861 Szymanski M.R., Fiebach A.R., Tratschin J.D., Gut M., Ramanujam V. M., Gottipati, K., Patel, P., Ye, M., Ruggli, N., and Choi, K. H. 2009. Zinc binding in pestivirus N(pro) is required for interferon regulatory factor 3 interaction and degradation. J Mol Biol 391: 438–449 Tarradas, J., de la Torre, M. E., Rosell, R., Perez, L. J., Pujols, J., Munoz, M., Munoz, I., Munoz, S., Abad, X., Domingo, M., Fraile, L. and Ganges, L. 2014. The impact of CSFV in the immune response to control infection. Vir. Res. 185: 82–91

Tasaka, M., N. Sakamoto, Y. Itakura, M. Nakagawa, Y. Itsui, Y. Sekine-Osajima, Y. Nishimura-Sakurai, C. H. Chen, M. Yoneyama, T. Fujita, T. Wakita, S. Maekawa, N. Enomoto, and M. Watanabe. 2007. Hepatitis C virus non-structural proteins responsible for suppression of the RIG-I/Cardif-induced interferon response. J. Gen. Virol. 88: 3323-3333.

Tautz, N., Elbers, K., Stoll, D., Meyers, G. and Thiel, H. J. 1997. Serine protease of pestiviruses: determination of cleavage sites. J. Virol. 71: 5415-5422.

Tellinghuisen, T. L., Marcotrigiano, J., Gorbalenya, A. E., and Rice, C. M. 2004. The NS5A protein of hepatitis C virus is a zinc metalloprotein. J. Biol. Chem. 279: 48576-48587.

Tellinghuisen, T. L., Paulson, M. S., and Rice, C. M. 2006. The NS5A protein of bovine viral diarrhea virus contains an essential zinc-binding site similar to the hepatitis C virus NS5A protein. J. Virol. 80: 7450-7458.

Terpstra, C. and Wensvoort, G. 1988. The protective value of vaccine-induced neutralizing antibody titres in swine fever. Vet. Microbiol. 16: 123–128.

133

Terpstra, C., Woortmeyer, R. and Bartelein, S. J. 1990. Development and properties of a cell culture produced vaccine for hog cholera based on the Chinese strain. Dtsch Tierarztl Wochenschr. 97: 77-79.

Tews B. A., Schurmann E. M., Meyers, G. 2009. Mutation of cysteine 171 of pestivirus E rns RNase prevents homodimer formation and leads to attenuation of classical swine fever virus. J. Virol. 83:4823–4834

Thiel, H. J., Stark, R., Weiland, E., Rumenapf, T. and Meyers, G. 1991. Hog cholera virus: molecular composition of virions from a pestivirus. J. Virol. 65: 4705–4712

Thiel, H. J., Plagemann, P. G. M., and Moennig, V. 1996. Pestiviruses, p.1059-1073. In B. N. Fields, D. M. Knipe, and P. M. Howley (ed), Fields Virology, 3rd Edition. Vol. 1. Lippincott- Raven Publishers, Philadelphia, PA.

Thompson, A.A., Zou, A., Yan, J., Duggal, R., Hao, W., Molina, D., Cronin, C.N., and Wells, P. A. 2009. Biochemical characterization of recombinant hepatitis C virus nonstructural protein 4B: evidence for ATP/GTP hydrolysis and adenylate kinase activity. Biochem. 48: 906– 916.

Tomoiu, A., Gravel, A., Tanguay, R. M., and Flamand, L. 2006. Functional interaction between human herpesvirus 6 immediate-early 2 protein and ubiquitin-conjugating enzyme 9 in the absence of sumoylation. J. Virol. 80: 10218–10228.

Tratschin, J. D., Moser, C., Ruggli, N. and Hofmann, M. A. 1998. Classical swine fever virus leader proteinase Npro is not required for viral replication in cell culture. J. Virol. 72: 7681-7684. van Gennip, H. G., van Rijn, P. A., Widjojoatmodjo, M. N. and Moormann R. J. 1999. Recovery of infectious classical swine fever virus (CSFV) from full-length genomic cDNA clones by a swine kidney cell line expressing bacteriophage T7 RNA polymerase. J. Virol. Methods. 78: 117-128. van Gennip H.G., van Rijn P.A., Widjojoatmodjo M.N., de Smit A.J., and Moormann R.J. 2000. Chimeric classical swine fever viruses containing envelope protein E(RNS) or E2 of bovine viral diarrhoea virus protect pigs against challenge with CSFV and induce a distinguishable antibody response. Vaccine. 19: 447-59. van Gennip, H. G. P., van Rijn, P. A., Widjojoatmodjo, M. N., de Smit, A. J. and Moormann, R. J. M. 2001. Chimeric classical swine fever viruses containing envelope protein Erns or E2 of bovine viral diarrhea virus protect pigs against challenge with CSFV and induce a distinguishable antibody response. Vaccine. 19: 447-459. von Heijne,G. 1990. The signal peptide. J. Membr. Biol. 115: 195–201. van Oirschot, J. T. 1986. Hog Cholera, p. 289. In B. S. A. D. Leman, R. D. Glock, W. L.

134

Mengeling, R. H. C. Penny, and E. Scholl (ed.), Diseases Of Swine, Sixth ed. Iowa State University Press, Ames.

Van Oirschot , J. T. and Terpstra. C. 1989 Hog cholera virus, p113-130. In: Pensaert, M. B. (ed.),Virus Infections of Porcines, Elsevier Science Publishers, B. V. Amsterdam. van Oirschot, J. T. 2003. Vaccinology of classical swine fever: from lab to field. Vet Microbiol. 96:367–384. van Rijn, P. A., van Gennip, H. G., de Meijer, E. J., and Moormann, R. J. 1993. Epitope mapping of envelope glycoprotein E1 of hog cholera virus strain Brescia. J. Gen.Virol. 74: 2053–2060. van Rijn, P. A., Miedema, G. K. W., Wensvoort, G., van Gennip, H. G. P., and Moormann, R. J. M. 1994. Antigenic structure of envelope glycoprotein E1 of hog cholera virus. J. Virol. 68: 3934–3942.

Van Rijn, P. A., Bossers, A., Wensvoort, G., and Moormann, R. J. 1996. Classical swine fever virus (CSFV) envelope glycoprotein E2 containing one structural antigenic unit protects pigs from lethal CSFV challenge. J. Gen. Virol. 77: 2737 - 2745. van Zijl, M., Wensvoort, G., de Kluyver, E., Hulst, M., van der Gulden, H., Geilkens, A., Berns, A. and Moormann, R. 1991. Live attenuated pseudorabies virus expressing envelope glycoprotein E1 of hog cholera virus protects swine against both pseudorabies and hog cholera. J. Virol. 65: 2761–2765.

Wang, Z., Nie, Y., Wang, P., Ding, M. and Deng, H. 2004 Characterization of classical swine fever virus entry by using pseudotyped viruses: E1 and E2 are sufficient to mediate viral entry. Virol. 330: 332–341.

Warrener, P. and Collett, M. S. 1995. Pestivirus NS3 (p80) protein possesses RNA helicase activity. J. Virol. 69: 1720–1726.

Watanabe, T., Wang, S., Noritake, J., Sato, K., Fukata, M., Takefuji, M., Nakagawa, M., Izumi, N., Akiyama, T., and Kaibuchi, K. 2004. Interaction with IQGAP1 links APC to Rac1, Cdc42, and actin filaments during cell polarization and migration. Dev. Cell 7: 871–883.

Wei, X., Decker, J. M., Wang, S., Hui, H., Kappes, J. C., Wu, X., Salazar-Gonzalez, J. F., Salazar, M. G., Kilby, J. M., Saag, M. S., Komarova, N. L., Nowak, M. A., Hahn, B. H., Kwong, P. D., and Shaw, G. M. 2003. Antibody neutralization and escape by HIV-1. Nature 422: 307–312.

Weiland, E., Stark, R., Haas, B., Rumenapf, T., Meyers, G. and Thiel, H. J. 1990. Pestivirus glycoprotein which induces neutralizing antibodies forms part of a disulfide-linked heterodimer. J. Virol. 64: 3563–3569

135

Weiland, E., Ahl, R., Stark, R., Weiland, F., and Thiel, H. J. 1992. A second envelope glycoprotein mediates neutralization of a pestivirus, hog cholera virus. J. Virol. 66: 3677–3682.

Weiland, F., Weiland, E., Unger, G., Saalmuller, A., and Thiel, H. J. 1999. Localization of pestiviral envelope proteins E(rns) and E2 at the cell surface and on isolated particles. J. Gen. Virol. 80: 1157–1165.

Weiskircher, E., Aligo, J., Ning, G., and Konan, K. V. 2009. Bovine viral diarrhea virus NS4B protein is an integral membrane protein associated with Golgi markers and rearranged host membranes. Virol. J. 6: 185. http://dx.doi.org/10.1186/1743-422X-6-185.

Westaway, E. G. 1987. Flavivirus replication strategy. Adv. Vir. Res. 33: 45-90.

Wengler, G., Bradley, D. W., Collett, M. S., Heinz, F. X., Schlesinger, R. W. and Strauss, J. H. 1995. Flaviviridae. In: F.A. Murphy, C. M. Fauquet, D. H. L. Bishop, S. A. Ghabrial, A. W. Jarvis, G. P. Marteli, M. A. Mayo, M.D. Summers (eds.), Virus Taxonomy, Sixth Report of the International Committee on Taxonomy of Viruses. Pp. 415-417. New York: Springer.

Wensvoort, G., Terpstra, C., and Bloemraad, M. 1986. Detection of antibodies against African swine fever virus using infected monolayers and monoclonal antibodies. Vet Rec. 28: 536-539.

Wensvoort G. 1989. Topographical and functional mapping of epitopes on hog cholera virus with monoclonal antibodies. J. Gen. Virol. 70: 2865–2876.

Wensvoort G., Boonstra J., and Bodzinga B. G. 1990. Immunoaffinity purification and characterization of the envelope protein E1 of hog cholera virus. J. Gen. Virol. 71:531–540

Widjojoatmodjo, M. N., van Gennip, H. G., Bouma, A., van Rijn, P. A., and Moormann, R. J. 2000. Classical swine fever virus E(rns) deletion mutants: Trans-complementation and potential use as nontransmissible, modified, live-attenuated marker vaccines. J. Virol. 74: 2973– 2980.

Wilson, M. E. 2003. The traveler and emerging infections: sentinel, courier, transmitter. J. Appl. Microbiol. 94: 1S–11S.

Wiskerchen, M. and Collett, M. S. 1991. Pestivirus gene expression: protein p80 of bovine viral diarrhea virus is a proteinase involved in polyprotein processing. Virol. 184: 341-50.

Xiao, M., Gao, J., Wang, W., Wang, Y., Chen, J., Chen, J. and Li, B. 2004. Specific interaction between the classical swine fever virus NS5B protein and the viral genome. Eur. J. Biochem. 271: 3888-3896.

Xiao, M., Wang, Y., Zhu, Z., Yu, J., Wan, L., and Chen, J. 2009. Influence of NS5A protein of classical swine fever virus (CSFV) on CSFV internal ribosome entry site-dependent translation. J. Gen. Virol. 90: 2923–2928.

136

Xu, J., Mendez, E., Caron, P. R., Lin, C., Murcko, M. A., Collett, M. S. and Rice, C. M. 1997. Bovine viral diarrhea virus NS3 serine proteinase: polyprotein cleavage sites, cofactor requirements, and molecular model of an enzyme essential for pestivirus replication. J. Virol. 71: 5312-5322.

Yamashita, T., Kaneko, S., Shirota, Y., Qin, W., Nomura, T., Kobayashi, K., and Murakami, S. 1998. RNA-dependent RNA polymerase activity of the soluble recombinant Hepatitis C Virus NS5B protein truncated at the C-terminal region. J. Biol. Chem. 273: 15479– 15486.

Yu, H., Grassmann, C. W., and Behrens, S. E. 1999. Sequence and structural elements at the 30 terminus of bovine viral diarrhea virus genomic RNA: Functional role during RNA replication. J. Virol. 73: 3638–3648.

Yu, H., Isken, O., Grassmann, C. W., and Behrens, S. E. 2000. A stem-loop motif formed by the immediate 50 terminus of the bovine viral diarrhea virus genome modulates translation as well as replication of the viral RNA. J. Virol. 74: 5825–5835.

Yueh, A., Leung, J., Bhattacharyya, S., Perrone, L.A., de los Santos, K., Pu, S. Y., and Goff, S. P. 2006. Interaction of Moloney murine leukemia virus capsid with Ubc9 and PIASy mediates SUMO-1 addition required early in infection. J. Virol. 80: 342–352.

Zhang G., Flick-Smith, H., McCauley, J. W. 2003. Differences in membrane association and sub-cellular distribution between NS2-3 and NS3 of bovine viral diarrhoea virus. Virus Res. 97: 89-102.

Zsak, L., Lu., Z., Kutish G. F., Neilan, J. G. and Rock, D. L. 1996. An African swine fever virus virulence associated gene Nl-S with similarity to the herpes simplex virus ICP34.5 gene. J. Virol. 70: 8865-8871.

Zogg, T., Sponring, M., Schindler, S., Koll, M., Schneider, R., Brandstetter, H., and Auer, B. 2013. Crystal structures of the viral protease Npro imply distinct roles for the catalytic water in catalysis. Structure 21: 929–938.

Zuker, M. 1989. The use of dynamic programming algorithms in RNA secondary structure prediction. Mathematical Methods for DNA Sequences, pages 159-184.

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Figure Figure 6. and BIC viruses anti

143

FlagT4 10-5 108

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Figure 7. Daily107105 temperature of animal groups in minimum protective dose study

(A) Flag T4v 101061045 TCID

105103

F Temperature 104102

103101

Temperature F Temperature 102100

10199 10098 1 dpi 6 dpi 11 dpi 16 dpi 21 dpi 5 dpc 10 dpc 15 dpc 20 dpc 99 Days postFlagT4-inoculation/Days 10-4 post-challenge 4 (B) Flag T4v 10108 98TCID 1 dpi 6 dpi 11 dpi 16 dpi 21 dpi 5 dpc 10 dpc 15 dpc 20 dpc 107 Days post-inoculation/Days post-challenge 106 105

104

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101 FlagT4 10-3 108100 (C) Flag T4v 10107399 TCID

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104

103

Temperature F Temperature 102

101

100

98 Mock inoculated control (D) Mock 1 dpi 6 dpi 11 dpi 16 dpi 21 dpi 5 dpc 10 dpc 15 dpc 20 dpc 108 Days post-infection/Days post-challenge 107

106

105

104

103

Temperature F Temperature 102

101

100 144 Days post infection/Days post challenge 99

98 1 dpi 6 dpi 11 dpi 16 dpi 21 dpi 5 dpc 10 dpc 15 dpc 20 dpc Days post-inoculation/Days post-challenge

Figure 8. Depiction illustrating altered codon usage for Flag T4G (A). Nucleotide changes for Flag region in E1 glycoprotein (B). Nucleotide substitutions for T4 region in envelope protein E2.

145

Figure 9. Growth Curve of Flag T4Gv compared with Flag T4v and BICv

146

Figure 10. Antigenic profile of Flag T4Gv compared to parental BICv and stained by immunohistochemistry with mAbs WH174, WH303, or anti-Flag (A). Flag T4Gv and BICv plaques shown at 1× magnification (B). Flag T4Gv and BICv plaques shown at 100× microscopic magnification.

(A)

(B)

147

Figure 11. Viremia Flag T4Gv Protection Assessment Experiment.

148

Figure 12. Viremia of Flag T4GV Early Protection Study

149

Figure 13. Cytokine Quantification in Serum

IFNa (ng ml) IFNb (ng ml) IFNg (pg ml) 600 500 4 400 400 3 300 2 200 200 100 1 0 0 0 Control BICv Flg T4 G Flg T4 G Control BICv Flg T4 G Flg T4 G Control BICv Flg T4 G Flg T4 G Naive & BICv Naive & BICv Naive & BICv

IL−1a (pg ml) IL−1b (pg ml) TGFb (ng ml)

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IL - 8 (pg ml) MCP2 CCL8 (pg ml) MX - 1 (ng ml) 80 400 7.5 60 300 40 200 5.0 20 100 2.5 0 0 0.0 Control BICv Flg T4 G Flg T4 G Control BICv Flg T4 G Flg T4 G Control BICv Flg T4 G Flg T4 G Naive & BICv Naive & BICv Naive & BICv

OAS1 (ng ml) PKR EIF2AK2 (ng ml) VCAM - 1 CD106 (ng ml) 20 0.5 0.4 40 15 0.3 10 20 0.2 5 0.1 0 0 0.0 Control BICv Flg T4 G Flg T4 G Control BICv Flg T4 G Flg T4 G Control BICv Flg T4 G Flg T4 G Naive & BICv Naive & BICv Naive & BICv 150

Figure 14. qRT-PCR Cytokine expression (A). (B).

151

Table 1. Amino Acid and Nucleotide Changes Observed in Revertant Flag T4SPv.

Position Changed CSFV Genomic Area

Substitution residue 109 (A to V), nucleotide 699 (C to T) NPro

Deletion residues 688 – 706, nucleotides 2436 – 2489 E1

Substitution residue 850 (N to S), nucleotide 2932 (A to G) E2

Silent mutation residue 861, nucleotide 2944 (A to G) E2

Silent mutation residue 866, nucleotide 2960 (A to C) E2

Substitution residue 993 (A to E), nucleotide 3351 (C to A) E2

Silent mutation residue 1360, nucleotide 4507 (T to C) NS23

Silent mutation residue, nucleotide 8170 (A to G) NS4B

Silent mutation residue, nucleotide 10177 (A to G) NS5B

Deletion nucleotide C-12342 3’UTR

152

Table 2. Nucleotide Sequence of Primers used for Sequencing cDNA Infectious Clones (F designates forward primer; R designates reverse primer).

Primer Name Sequence (5' - 3') F18 GTA TAC GAG GTT AGT TCA F200 GTC GTC AGT AGT TCG ACG R245 GGT TAA GGT GTG TCT TGG R483 TGT GGG TGT ACC TCA CT F556 CAT CAA AAC AAC ACT GAA F808 CTG CTG AAG CTA GCC AA R945 ATC CTC TTT TGG GGC TAT F1024 GGA AGG AGT AAA ATA CCA R1114 CCA ATA GGG CTT TTT CTA F1270 CAG AAG CTT GCA TGG GAT R1282 GGA CTC CTT TGC ATA TTT F1414 TGA ATG GAA CAA ACA TGG F1645 AGA GGG CCC ATG TAA TTT F1843 AGA AGC AAA ACC TGG TT R1888 TGT ACC ATA TGT ACC CTA F2008 GAT TCT CCA TGA GAT GGG R2046 GGC GAA GTC AGA CAG AAC F2379 CCT CAT CTG CTT GAT AAA AG F2599 CTG CAT GGC AGG TTC CTT R2715 GTC ATC CCC CAT TTC CTC R2886 GGT CTT TAC CAC TTC TGT F3007 GGG GGG CAA TTG GAC ATG R3361 AGC TGT CCC TGG GCT CAT F3421 CGA GTA TCA GTA CTG GTT R3539 AGC GGC GAG TTG TTC TGT F3784 GGG TGG AAA GAT AGA TGG R3831 ACT ACT ATA ACT GTC AG F3997 GCT AAC CTG GAC CTA CAT F4198 GAA TCT GGA CAT AGC CGG R4408 ACC CCT TCA CCA GCT TG F4369 ACC AAC TTC AAG AGG GT R4567 CTC CTG CTA TTT CAT CTA F4600 AGG GAC CAA CTT CAT CTC R4707 CCT CAC TTT GTG TTT GAT F4801 TGG CTT AGT CAA GGC AGC F4984 GGA TCA ATC AGA AGG GCC R5069 ACC AGG AGC ATC TTG ACT F5234 TTC GGT GTT ATG CCA AGG R5337 AGT GAC ATG GTC CAC TGA F5464 AGT TAA AAC TGA CTC CGG F5662 TGA GGC ATC AAG TGG AAG R5720 ACT GTT TGT ATT CCA CTC F5875 GTC ATA GAA GAG ATA GG F6062 CAG ATG CCA CAA CCT AAG 153

Primer Name Sequence (5' - 3') F6268 AGC CCC AGA AGT GAT GAA R6310 CCT CTA CTG GTA TCT TTA F6451 GGA GGA TCC ATC TAA CCT F6601 AAG ATG CCC TTC ATA GT R6636 CCT TCT CTG GGC TTG TTC F6787 GAT AAA CAT CAC CAA ATC R6915 TTC TGG GTG GTC AGT CCT F7039 TAA CTA TAC CTT CCT CAA R7083 AAG TCC TCA TCC TCT GT F7177 TGG TAG AGC ACT AAA ACA R7302 GTG GTG TCT TCC AAC CT F7402 TCA GGG GGA TGT GCA GAG F7588 ATA TGG GTT GTG GGG GAC R7713 TAA ACG ACC AAG TCT GT F7762 CCT CAG TTC CCA GGA GA R7935 ATG ACA ACG CTC TCC AA F8017 AAG CGA TGG TTT GCT AGG F8210 GAT GAA TTA GTC AAG GAG R8237 GTG TGC ACT GCT TCA AAC F8463 TCC TGT ATA AGT TCC GTG R8538 CCC GAG TCT GTC ATC CGT F8605 AAA TGC CCC TGT GGT TA R8751 TTC CAT TCT TAT CAC TGG R8824 ACT GGA TTT CAA CAA CAG R8872 TCT TGA GCA CCC TGA CCA F8965 CTG ATA GAG AGG GAC TG R9063 TTC TTG TGT ACC AAT TC F9180 ACC AGC CTT CGG GGA GAA F9422 CTG CAC GAA GCC ATA CAA R9467 ACT CTA TTA GAG GTG GCT F9659 ACC CTA GAG GCA TTA AGT R9682 CTT CTG CCT TGG TTA TGT F9831 GGG ATA TAG CTA CTA TCA R9853 TTT GTG GAG TCT GTA GCC F10039 GGT CTC TGC TTA CCA ACT R10143 GTG TTC CTC TAC CAA CTC F10167 TAG AGG AAC ACA AGA TGA R10272 CAG TTG CTC TGC CAC CTT F10412 GAC ACA ACC AAC TTC CAC R10500 CTC TAA CTC GGG TCG TTT F10658 GAG ATT ATT GAC AAT CTG R10774 TTG CTT CAG GGT ATT GTA R10931 TCG AAG CTC ACT GCC ACT F11080 TAT GTG AGA AGT ACC CGT F11275 GAT GGT TTC CTG ATC AC F11416 GTT TTG CTC CCA TAC ACC

154

Primer Name Sequence (5' - 3') F11650 GAA ACC AGG GAA GTC AAC R11727 CAG CTT CTC GAA GCT TGT F11824 ACT ACA AGA CTG TGT CAA R12222 TGA GTG TAG TGT GGT AAC F12058 GAT GAT GAC CCT GAT AGG R12307 GAG CCA TGG CCC GGG CCG TTA GAA ATT A

155

Table 3. Nucleotide Sequence of Primers used for Evaluation of Revertant Flag T4SPV (F designates forward primer; R designates reverse primer).

Primer Name Sequence (5' - 3')

F699 CTAGAGTTTTTTGACGAAGTACAGTTTTGTGAGGTGACC

R699 GGTCACCTCACAAAACTGTACTTCGTCAAAAAACTCTAG

F2932 GGTGTTATAGAGTGCACGTCATTTAGTATGGACACTCTG

R2932 CAGAGTGTCCATACTAAATGACGTGCACTCTATAACACC

F3551 TAAGGAGATCGTCTCTAGTGATGGACCTGTAAGGAAAACTTC

R3551 GAAGTTTTCCTTACAGGTCCATCACTAGAGACGATCTCCTTA

156

Table 4. Nucleotide sequence of primers for Flag T4SPv PCR amplification (F designates forward primer; R designates reverse primer).

Primer Name Sequence (5' - 3')

F18 GTA TAC GAG GTT AGT TCA

R3361 AGC TGT CCC TGG GCT CAT

F3007 GGG GGG CAA TTG GAC ATG

R6310 CCT CTA CTG GTA TCT TTA

F6062 CAG ATG CCA CAA CCT AAG

R9063 TTC TTG TGT ACC AAT TC

F8965 CTG ATA GAG AGG GAC TG

R12307 GAG CCA TGG CCC GGG CCG TTA GAA ATT A

157

Table 5. Determination of Minimum Protective Dose Experiment - Swine Survival and Fever Response Following Dose Ranging Infections with Flag T4v and Lethal Challenge 21 dpi.

(a) One animal dye at day 6 post FlagT4 inoculation. Necropsy showed no CSFV/ ASFV/FMDV related symptoms. (b) Two animals were euthanized at days 9 and 15 post challenge presenting CSFV related symptoms. (c) Two animals developed fever at day 6 post challenge. (d) Two animals were euthanized at days 8 and 13 post challenge presenting CSFV related symptoms. (e) Two animals developed fever at days 3 and 5 post challenge. (f) Values are calculated based only in data from the two affected animals. All other animals in the group were clinically normal. (g) Animals were euthanized at days 7 (two animals), 8 (one animal), 9 (two animals), 10 (two animals) and 11 (three animals) post challenge presenting CSFV related symptoms. (h) Challenge was performed by IN inoculation of 105.13 TCID50 of BICv. (i) Back-titration indicate that each group received approximately one log less of FlagT4v than intended

158

Table 6. Survival and Fever Response in Pigs Infected with CSFV Chimeric Viruses Representing Mutations in Flag T4SPv

Fever

No. of Mean time to No. of days to Duration Virus survivors/total death onset (days + SD) No. (days + SD ) (days + SD)

BICv 0/4 9.76 (0.95) 4 (0.0) .75 (0.5)

FlagT4 699v 4/4 - - -

FlagT4 2932v 4/4 - - -

FlagT4 3351v 4/4 - - -

T4 699v 4/4 - - -

T4 2932v 0/4 11.25 (0.96) 3.5 (0.5) 7.75 (2.1)

T4 3351v 4/4 - - -

159

Table 7. Nucleotide Sequence of Primers used for the Production of Flag T4Gv (F designates forward primer; R designates reverse primer).

Primer Name Sequence (5' - 3')

F T4G GACGGGTGTTATAGAGTGCAGCAGTTTCGGAATGGATACTCTGAGA

R T4G TCTCAGAGTATCCATTCCGAAACTGCTGCACTCTATAACACCCGTC

F Flag1 GTTACTGGTAACTGGGGCACCAGTGAGTTGCACCCACCTTGGCAGATTACAAG

R Flag1 CTTGTAATCTGCCAAGGTGGGTGCAACTCACTGGTGCCCCAGTTACCAGTAAC

F Flag2 CACCATGTAGTTGCACCCACTTAGCCGCGGCCGACTATAAGGATGACGACGATAAGCA

R Flag2 TGCTTATCGTCGTCATCCTTATAGTCGGCCGCGGCTAAGTGGGTGCAACTACATGGTG

F Flag3 CCCACTTGACCGCGGCCGACTATAAAGACGATGATGACAAACAAGGCCGGCTAGC

R Flag3 GCTAGCCGGCCTTGTTTGTCATCATCGTCTTTATAGTCGGCCGCGGTCAAGTGGG

160

Table 8. Swine survival and fever response in Flag T4Gv infected animals after challenge with parental virulent BICv.

Fever

Mean time No. of days Duration Maximum No. of Challenge Group to death to onset No. of days daily temp, survivors/total (+ SD) (+ SD) (+ SD) °F (+ SD)

Mock 0/4 10.25 (0.96) 4.25 (0.96) 6 (1.41) 106.1 (0.31)

Flag T4Gv 7 DPI 4/4 0 0 0 103.2 (0.38)

Flag T4Gv 28 DPI 4/4 0 0 0 103.1 (0.78)

161

Table 9. Swine Survival and Fever Response in Flag T4Gv Infected Animals Challenged with BICv at Different Time Points Post Infection.

Fever

Mean time No. of days Duration Maximum No. of Challenge Group to death to onset No. of days daily temp, survivors/total (+ SD) (+ SD) (+ SD) °F (+ SD)

Mock 0/5 9 (2.45) 5 (0) 3.8 (2.17) 105.6 (0.42)

FlagT4Gv 7 DPI 5/5 - - - 103.3 (0.31)

FlagT4Gv 5 DPI 5/5 - - - 103.4 (0.65)

FlagT4Gv 3 DPI 5/5 - - - 103.4 (0.58)

FlagT4Gv 2 DPI 3/5 13 (1.41) (1) 6.8 (1.26) (1) 4.5 (2.36) (1) 105.5 (1.66)(1)

FlagT4Gv 1 DPI 0/5 10 (2.82) 5 (0) 5 (2.82) 106.4 (1.22)

(1) Data are based in 2 out 5 animals presenting severe disease symptoms and ultimately euthanized at days 12 ad 14 days post challenge, and 1 out 5 animals presenting transitory rise of body temperature without any other CSF-related symptom. The other two animals remained clinically normal with maximum average daily body temperature of 103.6.

162

Table 10. Primer Sets for qRT-PCR Cytokine Gene Expression Assay.

Genes Forward primer (5′ → 3′) Reverse primer (5′→ 3′)

Cytokines

IL-1α GTGCTCAAAACGAAGACGAACC CATATTGCCATGCTTTTCCCAGAA IL-1β AACGTGCAGTCTATGGAGT GAACACCACTTCTCTCTTCA IL-2 TCAACTCCTGCCACAATGTATAAGA CTTGAAGTAGGTGCACCGTTTG IL-4 GCCGGGCCTCGACTGT TCCGCTCAGGAGGCTCTTC IL-5 CGTTAGTGCCATTGCTGTAGAAA CAAGTTCCCATCGCCTATCAG IL-6 CTGGCAGAAAACAACCTGAACC TGATTCTCATCAAGCAGGTCTCC IL-7 GAGTGACTATGGGCGGTGAGA GCGGGCGTGGTCATGA IL-10 CGGCGCTGTCATCAATTTCTG CCCCTCTCTTGGAGCTTGCTA IL-12p35 CGTGCCTCGGGCAATTATA CGCAGGTGAGGTCGCTAGTT IL-12p40 AACTCTTCACGGACCAAATCTCA GGTCCCGGGCTTGCA IL-13 CCTGGAATCCCTCATCAACATC AGGGCGCTCAGCATCCT IL-15 GCTGTATCAGTGCAGGTCTTCCT TTTCAGTATACAATGTGGCATCCA IL-16 CGAAGACCCAGGTGCAAATAGT CCGAAAGGTTGAGCGAGAAG TNF-α AACCTCAGATAAGCCCGTCG ACCACCAGCTGGTTGTCTTT TGF-β1 CCTGCAAGACCATCGACATG GCCGAAGCTTGGACAGAATC TGF-β2 TGTGTGCTGAGCGCTTTTCT GAGCGTGCTGCAGGTAGACA NCP-1 CGCCATCTCCTCCTGTACCA GCAGTGACAATGGCAGCAAT Chemokines

IL-8 AAGCTTGTCAATGGAAAAGAG CTGTTGTTGTTGCTTCTCAG AMCF-1 GCTCGTGTCAACATGACTTCCA GCCTCACAGAGAGCTGCAGAA AMCF-2 CCACACCCGGGATTCATC GGCTATCACTTCTGCCTTGGA MCP-1 GCAGCAAGTGTCCTAAAGAAGCA GCTTGGGTTCTGCACAGATCT MCP-2 AAGACCAAAGCCGACAAGGA TCATGGAATTCTGGACCCACTT RANTES AGCATCAGCCTCCCCATATG TTGCTGCTGGTGTAGAAATATTCC

______

163

Genes Forward primer (5′ → 3′) Reverse primer (5′→ 3′)

Toxicity response

iNOS GCGATGGGAAGCACGACTT CATCTGGTAGCCGGCATACC

IL-1β converting enzyme

ICE GCCAAGAGGGAGCCTCAAG CTCTGCTGACTTTTCTTTCCATAGC Adhesion molecules

ICAM CACAGGCCGCCACTAACAA GGTTCCATTGATCCAGGTCTTG VCAM GCTCCCAGGGATACGACCAT ACTAGAGCAGGTCATGTTCACAGAA Antiviral effect

PKR AAAGCGGACAAGTCGAAAGG TCCACTTCATTTCCATAGTCTTCTG OAS GAGCTGCAGCGAGACTTCCT TGCTTGACAAGGCGGATGA Mx1 GGCGTGGGAATCAGTCATG AGGAAGGTCTATGAGGGTCAGATCT Interferons and interferon regulatory factors

IFN-α TCTCATGCACCAGAGCCA CCTGGACCACAGAAGGGA IFN-β AGTGCATCCTCCAAATCGCT GCTCATGGAAAGAGCTGTGGT IFN-γ CGATCCTAAAGGACTATTTTAATG TTTTGTCACTCTCCTCTTTCCAAT IRF-1 GCACCAGCGACCTGTACAACT TCCTCATCTGTTGCAGCTTCA IRF-3 CGCTTCTTGCCCCAACCT TCCCACTCGTCGTCATTCG IRF-6 GTGTACTGGTCTGGGCCATGT GACCTTCTTTTGCCTCTCAATCA IRF-7 CGCCTCCTGGAAAACCAA CCCTGAGTTGTCCTGCAACA IRF-9 GGCCCTGCCATCTGGAA CAGGAACCTCCTCAAACTCAGTACT

Complement regulatory proteins

CD46 GTGTCGTTGGCCCTAGTGTTC GGCCGCATGCTTTCAAACT

______

164

Genes Forward primer (5′ → 3′) Reverse primer (5′→ 3′) Cytokine receptors

IL-1R TTGCTTTGCAGGTAATCATCAGA TGCTGAACAAGGACACCACAGT IL-2R-α GTATGCGGGCTGGATGCT TGCCTAAGCTGTTGCTATTCCAA IL-6R GGTACCATTGCCCACATTCC GAACCTTAAGATGATGCCAATGC IL-8R CATCTTCCGTGAAGCCTACCA TTGTATTGGCGCCCAGATC IL-12R CTGACTCAAAGCCGGGAAAC ATCCATGTTGGAGGGTAAGTAACC TNF-R CCGCTGGTGATCTTCTTTGG CGTTGGTAGCGGCAAGCT TGF-β3R TTCTCCACCGATTTTTCATGGT AGCAGAGCTCCGATCACAAAC DAP-12 GAGCCCAAATCAGGACAGTCA GTATCCTGGTGGGAGCAGAAAG NKG2D AATCACAAGCAAATGTGGAGAAAA AAACGGATCCCCATAGCTATAGC

Toll-like receptors

TLR-1 TCCTGATCTTGCTGGATTCCA TCCTTGGGCCATTCCAAA TLR-2 GGCAAGTGGATTATTGACAACATC ACCACTCGCTCTTCACAAAGTTC TLR-3 CAAAACCAGCAACACGACTTTC AATCATTACCAATCACACTTAAGCT TLR-4 CCTCGGAGGCTGCTCTGA GCGGATGCGAGGAATCAT TLR-5 TGCTAGATGTTTCTGGCAATGG GGCGAGAACCAAGGAGGAA TLR-6 TCTCACAATCAGTTGCAGACAATCT CATATGGGCAGGGCTTCAAA TLR-7 CAAGCACTTGACAGCGATTCC GGAAGGAGGCTGGAGTGATG TLR-8 AAGGGCTGCAAAATCTGACTAAAA AAACGCCCCATCTGTAATAGTCAT TLR-9 TGTCCAGCCTACGAACTCTCAA GTTGGGCTCGATGGTCATGT TLR-10 ACTGCATGGAGAAAAGCTCTAAGTC GAGGTGGTGGTGGCCAAAG

IL, interleukin; TNF, tumor necrosis factor; TGF, transforming growth factor; NCP, neutrophil chemotactic protein; AMCF, alveolar macrophage chemotactic factor; MCP, monocyte chemotactic protein; ICAM, intercellular adhesion molecule; VCAM, vascular cell adhesion molecule; PKR, protein kinase R; OAS, 2′-5′-oligoadenylate synthetase; Mx, interferon-inducible GTPases; IFN, interferon; IRF, interferon regulatory factor; CD, cluster of differentiation; DAP, transmembrane adaptor protein; NKG, natural killer cell receptor; TLR, toll-like receptor.

165